FRESHWATER MOLLUSKBIOLOGY ANDCONSERVATIONTHE JOURNAL OF THE FRESHWATERMOLLUSK CONSERVATION SOCIETY

VOLUME 19 NUMBER 1 MAY 2016

Pages 1-21A National Strategy for the Conservation of Native Freshwater MollusksFreshwater Mollusk Conservation Society

Pages 22-28Investigation of Freshwater Mussel Glochidia Presence on Asian Carp and Native Fishes of the Illinois RiverAndrea K. Fritts, Alison P. Stodola, Sarah A. Douglass and Rachel M. Vinsel

Pages 29-35Toxicity of Sodium Dodecyl Sulfate to Federally Threatened and Petitioned Freshwater Mollusk SpeciesKesley J. Gibson, Jonathan M. Miller, Paul D. Johnson and Paul M. Stewart

Freshwater Mollusk Biology and Conservation

©2016ISSN 2472-2944

Editorial Board

CO-EDITORSGregory Cope, North Carolina State University

Wendell Haag, U.S. Department of Agriculture Forest ServiceTom Watters, The Ohio State University

EDITORIAL REVIEW BOARDConservation

Jess Jones, U.S. Fish & Wildlife Service / Virginia Tech University

EcologyRyan Evans, Kentucky Department of Environmental Protection, Division of Water

Michael Gangloff, Appalachian State UniversityCaryn Vaughn, Oklahoma Biological Survey, University of Oklahoma

Freshwater GastropodsPaul Johnson, Alabama Aquatic Biodiversity Center

Jeff Powell, U.S. Fish & Wildlife Service, Daphne, AlabamaJeremy Tiemann, Illinois Natural History Survey

Reproductive BiologyJeff Garner, Alabama Division of Wildlife and Freshwater Fisheries

Mark Hove, Macalester College / University of Minnesota

Survey/MethodsHeidi Dunn, Ecological Specialists, Inc.

Patricia Morrison, U.S. Fish & Wildlife Service Ohio River Islands RefugeDavid Strayer, Cary Institute of Ecosystem Studies

Greg Zimmerman, Enviroscience, Inc.

Systematics/PhylogeneticsArthur Bogan, North Carolina State Museum of Natural Sciences

Daniel Graf, University of Wisconsin-Stevens PointRandy Hoeh, Kent State University

ToxicologyThomas Augspurger, U.S. Fish & Wildlife Service, Raleigh, North Carolina

Robert Bringolf, University of GeorgiaJohn Van Hassel, American Electric Power

Teresa Newton, USGS, Upper Midwest Environmental Sciences Center

Freshwater Mollusk Biology and Conservation 19:1–21, 2016

� Freshwater Mollusk Conservation Society 2016

ARTICLE

A NATIONAL STRATEGY FOR THE CONSERVATION OFNATIVE FRESHWATER MOLLUSKS*

Freshwater Mollusk Conservation Society**,1

1417 Hoff Industrial Dr., O’Fallon, MO 63366 USA

ABSTRACT

In 1998, a strategy document outlining the most pressing issues facing the conservation offreshwater mussels was published (NNMCC 1998). Beginning in 2011, the Freshwater MolluskConservation Society began updating that strategy, including broadening the scope to includefreshwater snails. Although both strategy documents contained 10 issues that were deemed prioritiesfor mollusk conservation, the identity of these issues has changed. For example, some issues (e.g.,controlling dreissenid mussels, technology to propagate and reintroduce mussels, techniques totranslocate adult mussels) were identified in the 1998 strategy, but are less prominent in the revisedstrategy, due to changing priorities and progress that has been made on these issues. In contrast, someissues (e.g., biology, ecology, habitat, funding) remain prominent concerns facing mollusk conservationin both strategies. In addition, the revised strategy contains a few issues (e.g., newly emerging stressors,education and training of the next generation of resource managers) that were not explicitly present inthe 1998 strategy. The revised strategy states that to effectively conserve freshwater mollusks, we needto (1) increase knowledge of their distribution and taxonomy at multiple scales; (2) address the impactsof past, ongoing, and newly emerging stressors; (3) understand and conserve the quantity and quality ofsuitable habitat; (4) understand their ecology at the individual, population, and community levels; (5)restore abundant and diverse populations until they are self-sustaining; (6) identify the ecosystemservices provided by mollusks and their habitats; (7) strengthen advocacy for mollusks and theirhabitats; (8) educate and train the conservation community and future generations of resourcemanagers and researchers; (9) seek long-term funding to support conservation efforts; and (10)coordinate development of an updated and revised strategy every 15 years. Collectively addressingthese issues should strengthen conservation efforts for North American freshwater mollusks.

KEY WORDS - freshwater mollusks, conservation, management, strategy, snails, mussels

INTRODUCTION

In 1995, federal, state, academic, and private sector

managers and researchers concerned with the decline of

freshwater mussels (order Unionoida, families Unionidae and

Margaritiferidae) in North America formed the National

Native Mollusk Conservation Committee (NNMCC) to

discuss the conservation status of this imperiled fauna.

Realizing the scope and immediacy of the issues associated

with mussel imperilment, the NNMCC decided that a

nationwide coordinated effort was needed to stem population

declines and extinctions. They produced a document to

address the conservation needs of mussels and drafts of that

document—the National Strategy for the Conservation of

Native Freshwater Mussels (hereafter referred to as the

National Strategy)—were presented at two North American

1

*Article Citation: FMCS (Freshwater Mollusk Conservation Society).

2016. A national strategy for the conservation of native freshwater

mollusks. Freshwater Mollusk Biology and Conservation 19: 1-21.**Corresponding Author: tnewton@usgs.gov

1Contributing authors listed alphabetically, as follows: Megan Bradley,

U.S. Fish and Wildlife Service; Robert S. Butler, U.S. Fish and Wildlife

Service; Heidi L. Dunn, Ecological Specialists, Inc.; Catherine Gatenby,

U.S. Fish and Wildlife Service; Patricia A. Morrison, U.S. Fish and

Wildlife Service; Teresa J. Newton, U.S. Geological Survey; Matthew

Patterson, U.S. Fish and Wildlife Service; Kathryn E. Perez, University of

Texas Rio Grande Valley; Caryn C. Vaughn, Oklahoma Biological Survey

and Department of Biology, University of Oklahoma; Daelyn A.

Woolnough, Central Michigan University, Biology Department and

Institute for Great Lakes Research; Steve J. Zigler, U.S. Geological Survey

mussel symposia (in 1995 and 1997). In 1998, the National

Strategy was published in the Journal of Shellfish Research(NNMCC 1998). This document has been used to prioritize

research and management actions related to mussel conserva-

tion. In addition, the National Strategy has helped organize

activities conducted by the Freshwater Mollusk Conservation

Society (FMCS)—the organization that evolved from the

NNMCC—that is dedicated to the conservation and advocacy

of freshwater mollusks. For brevity, we refer to native

freshwater mussels (mussels) and gastropods (snails) simply

as mollusks.

The FMCS was initiated as a grass-roots effort by

dedicated individuals among agencies and organizations,

academia and private citizens, and across states and countries.

The work of this society has resulted in the recruitment and

training of many scientists and managers who work at local,

state, regional, national, and international levels on mollusk

conservation. As a consequence, many published papers,

agency reports, and government documents have been

generated by this growing community. Our understanding of

mollusks and the ecological role they play in aquatic

ecosystems has grown exponentially as reflected in the peer-

reviewed literature (Figure 1). Communication and outreach

on mollusks has also increased the scope and breadth of

awareness of the conservation challenges facing managers and

researchers to wider audiences, including decision and policy

makers. As a result of enhanced partnerships, funding to

protect mollusks and their habitats has increased.

Haag and Williams (2014) assessed the state of progress

made under each of the 10 ‘‘problems’’ outlined in the 1998

National Strategy and suggested ways to improve conservation

implementation. After nearly two decades of progress and

change, it was clear that an updated document was needed to

address evolving conservation challenges, research needs, and

emerging threats for mollusks. In 2011, the FMCS formed a

committee to revise the National Strategy, and, in 2013, a list

of overarching issues in mollusk conservation and strategies to

address them was approved by the FMCS membership. Here,

we present those issues as an updated National Strategy

intended to identify research, monitoring, management, and

conservation actions needed to sustain and recover mollusks.

The issues are presented without prioritization because the

strategies within the issues are independent and may vary due

to numerous factors. This manuscript intends to give guidance

to our conservation partners, but is not intended to prioritize

action—that is a decision that should be made by local, state,

or regional partners based on regional and taxonomic

priorities, funding, expertise, etc.

The 1998 National Strategy was restricted to the

conservation of mussels; however, many of the same factors

that led to the imperilment of mussels has also affected snails.

Recent compilations indicate that both groups have a similar

high level of continental imperilment (~74%, J.D. Williams,

Florida Museum of Natural History, personal communication;

Johnson et al. 2013). In addition, snails are less studied than

mussels, with most species defined only in terms of their

taxonomy (see Graf 2001; Perez and Minton 2008). However,

there is a growing body of knowledge regarding their

distribution, ecology, and conservation status (Brown et al.

2008; Lysne et al. 2008; Johnson et al. 2013). We know that

both mussels and snails require healthy aquatic ecosystems,

but their specific life histories and ecological requirements can

be quite different. For example, snails often occur in different

habitats than mussels, have fundamentally different life

histories, have higher rates of endemism, often have highly

restricted distributions, and have dispersal mechanisms not

dependent on hosts. The diversity of the snail fauna in the U.S.

and Canada (16 families) implies a wide range of life history

traits and needs (Johnson et al. 2013). These requirements will

need to be considered, in addition to those for mussels, for

effective conservation of the mollusk fauna.

This National Strategy focuses largely on conservation

issues in the U.S. and Canada, although similar threats to

mollusks and their habitats are occurring worldwide (Lydeard

et al. 2004). We acknowledge that the factors affecting

mollusk communities vary in scope and intensity across

regions. In the interest of brevity, we do not address region-

specific issues or provide an exhaustive literature review for

the broad issues we do cover.

The goal of this National Strategy is to identify and

clarify issues and actions that are essential to conserving our

nation’s mollusk fauna and ensure that their ecological,

social, and economic values to society are maintained at

sustainable levels. This revised strategy does not address

fingernail clams (Sphaeriidae), largely because not enough

data or expertise is available to include them. This document

presents the 10 issues that are considered priorities for the

conservation of mollusks. For each issue, we state the overall

goal, list strategies needed to address the issue, provide an

Figure 1. Number of peer-reviewed scientific articles during 1990–2014 that

referenced freshwater mollusks as their topic. Data from Web of Science

(February 2015) and are per year (i.e., not cumulative).

2 FMCS 2016

overview of the topic, offer examples of successes since the

1998 National Strategy, identify research needs, and

recognize opportunities that will aid the conservation and

management of mollusks.

Issue 1 – Increase knowledge of the distribution andtaxonomy of mollusks at multiple scales over time and makethat information available.

Goal: Understand the status and trends of mollusk

populations to better manage and conserve species.

Strategies:

1. Continue to refine knowledge of systematics, taxonomy,

and genetic structure of species.

2. Update and maintain a database of the accepted scientific

nomenclature.

3. Use survey methods that provide data needed for trend

analyses.

4. Identify uniform data collection and reporting standards

that will support periodic status assessments.

5. Encourage reporting of distribution data.

6. Assess and publish the conservation status of mollusks

every 10 years.

Overview

Knowledge of the distribution of mollusks is lacking,

which hinders our ability to manage their populations. A key

difficulty in understanding the distribution of mollusks has

been poor taxonomy. Although considerable progress has been

made in taxonomy since 1998 (Bogan and Roe 2008; Johnson

et al. 2013), most species have not been studied using modern

techniques. Many waterbodies are virtually unsampled, and

even in well-studied regions, many drainages have not been

adequately surveyed in half a century or more. An update of

our approaches to taxonomy and generating and managing

distribution data is needed to make this information more

accurate and widely available.

Success stories in mollusk distributions

Numerous stream and lake surveys have been published

since 1998. Many of these studies have resulted in range

extensions, new drainage records, population trends, and

demographic data (e.g., Evans and Ray 2010; Haag and

Warren 2010; Zanatta et al. 2015). Compilation of distribu-

tional information for species status assessments, recovery

plans listed under the Endangered Species Act (ESA), and

other studies have contributed to a greater understanding of

mollusk distributions, which are critical for conservation

efforts (e.g., U.S. Fish and Wildlife Service [USFWS] 2004;

Butler 2007; Crabtree and Smith 2009). One example of a

broad-scale survey was conducted in Texas rivers in 2010

(Burlakova and Karatayev 2010; Burlakova et al. 2011). New

populations of endangered species and fewer populations of

other species were documented, leading to a more accurate

assessment of the global status of these species, including a

recommendation that they be listed as endangered by Texas.

Some advances have been made in applying molecular and

other data to the phylogenies of mollusks, thus facilitating a

better understanding of their distribution. A review of the

systematics of mussels was published in 2007 (Graf and

Cummings 2007) and a number of papers describing new

species or evolutionarily significant units have contributed to

their conservation (Serb et al. 2003; Bogan and Roe 2008;

Chong et al. 2008). Despite these efforts, the population

genetics of most mussel species remain completely unexplored.

Molecular phylogenies of some Physidae and Pleuroceridae

snails have been published (Lydeard et al. 1997; Holznagel and

Lydeard 2000; Minton and Lydeard 2003; Wethington and

Lydeard 2007), but recent data suggest mitochondrial molec-

ular data are unreliable for Pleuroceridae (Whelan and Strong

2016). Dozens of species of Hydrobiidae and Lithoglyphidae

snails have been described from the Southwestern and

Southeastern U.S. through the use of molecular data, electron

microscopy, and detailed anatomical review (Hershler et al.

2008; Hershler and Liu 2009, 2010). However, most snail

families remain unexamined using modern techniques. Data

from molecular studies have been used to strengthen

conservation efforts for some mussel (Jones et al. 2006; Jones

and Neves 2010; Jones et al. 2015) and snail (Wethington and

Guralnick 2004; Morningstar et al. 2014) species.

Several efforts to make mollusk information publicly

available have gained momentum. For example, papers

updating the continental status and summarizing the known

distributions of mussels and snails have been published

(Williams et al. 1993; Johnson et al. 2013). The Freshwater

Gastropods of North America (Dillon et al. 2013) contains

distribution and ecological information organized by geo-

graphic region. Keys to the snails of some U.S. states have

become available online (e.g., Thompson 2004; Perez and

Sandland 2015). NatureServe and the American Fisheries

Society databases remain the primary sources for distribution

information, though both have significant limitations (Wil-

liams et al. 1993; Johnson et al. 2013; NatureServe 2015).

Research to aid conservation and management

In addition to the basic need for alpha taxonomy, detailed

phylogenetic and population genetic studies are also needed.

Analyzing datasets across multiple sources (e.g., multiple

genes, microsatellites, morphology, ecological relationships)

will ensure that these data are accurate and representative of

natural populations. Collectively, these data may be important

in recognizing patterns of imperilment of species, species

groups, and genera. Determining the genetic structure of

healthy populations will serve as a benchmark for the levels of

genetic diversity and composition necessary for sustainability.

The last assessment on the names of mollusks was

conducted by Turgeon et al. (1998); this list needs immediate

revision. Taxonomic changes that have occurred among

mollusks need to be updated and these findings need to be

3NATIONAL STRATEGY FOR FRESHWATER MOLLUSKS

published every 10 years. Maintaining an ongoing, publicly-

available record of accepted names would be ideal (e.g.,

fishbase.org). Similarly, comprehensive conservation status

assessments for mussels and snails should be conducted every

10 years. The first snail assessment was published in 2013

(Johnson et al. 2013), and the second mussel assessment is in

progress (J.D. Williams, Florida Museum of Natural History,

personal communication). Ideally, future assessments will

have more spatially-explicit distribution data, such as major

river drainages, rather than simply states and provinces of

occurrence. Further, efforts should transition to up-to-date,

accessible online databases, rather than solely periodic

publications. Online museum databases are useful for

distribution records in some groups, (e.g., taxa with unique

shell morphologies), but are questionable for other mollusks

(especially snails), as no museum has comprehensively

revisited the current identifications of mollusk holdings.

Therefore, online distribution records are often incorrect,

compromising their usefulness (Graf 2013; Johnson et al.

2013). Obtaining funding for maintaining taxonomic and

distributional databases is a critical conservation need.

Standardized sampling methods should be used to assess

mollusks over space and time. Although guidelines for

standardized sampling methods exist for mussels (e.g., Strayer

and Smith 2003), a companion work for snails has not been

developed. Sampling methods will vary by objective and

system type (e.g., wadeable streams, large rivers, lakes), and

ideally contain a quantitative component so that results can be

statistically evaluated across species, localities, watersheds,

and time. In addition to the number of mollusks sampled alive,

data such as shell-only individuals, shell condition (i.e., fresh

dead or relic), sex ratio, length, and age may be useful for

assessing status and trends. Also, data on habitat conditions,

potential stressors, and characteristics of aquatic communities

(e.g., host presence) may improve understanding of status and

trend information on mollusks and inform conservation.

Opportunities for conservation and management

Inadequate taxonomy and poor understanding of species

distributions make species extinctions more likely and

challenge effective conservation and management (May

1990; Perez and Minton 2008). For example, a rare species

that is superficially similar to a more common relative may be

overlooked and conservation measures not taken because it is

unobserved when the rare species declines in abundance. If

species distributions are not well-documented, then we cannot

tell when populations are lost to human actions. The

decreasing costs and increasing availability of genomic tools

such as microsatellites and next-generation sequencing offer

managers and researchers an opportunity to capture needed

data on systematic relationships, taxonomy, and ultimately

species distributions by identifying all ecologically significant

units within a species. These new techniques offer great

promise for the conservation of mollusks.

Our understanding of the conservation status of mollusks

at the state level has increased substantially due to databases

maintained under their Heritage programs and Comprehen-

sive Wildlife Action Plans (CWAP), in addition to the

publication of faunal books (e.g., Williams et al. 2008;

Watters et al. 2009). These efforts have facilitated more

accurate continental scale faunal assessments. Unlike the first

mussel assessment, where nearly 5% of the fauna were of

undetermined status (Williams et al. 1993), the forthcoming

mussel update assesses status for all species (J.D. Williams,

Florida Museum of Natural History, personal communica-

tion). In contrast, a recent assessment of the conservation

status for snails indicated that the status of 4% of species was

undetermined (Johnson et al. 2013). We need to update the

status of the 4% of the snail taxa that are still undetermined

and develop conservation plans that cover entire faunal

regions (e.g., Mobile, Ohio, Mississippi). Improving our

knowledge of taxonomy, life history traits, distribution,

dispersal routes, and population connectivity will help

managers prioritize imperiled species for conservation and

increase the likelihood that more faunas can be conserved.

This could lay the foundation for the partial restoration of

ecosystem services that mollusks provide to aquatic and

terrestrial organisms, ultimately including mankind (see Issue

6).

Issue 2 – Address the impacts of past, ongoing, and newlyemerging stressors on mollusks and their habitats.

Goal: Minimize threats to mollusks and their habitats.

Strategies:

1. Prepare white papers on known stressors (e.g., impound-

ments, dredging, contaminants, runoff, invasive species,

disease) describing the risk and magnitude of effects on

mollusks.

2. Describe the impacts of emerging stressors (e.g., climate

change, increased energy development, water use conflicts,

unregulated contaminants, hormone disruptors) and possi-

ble synergistic effects on mollusk populations.

3. Compile comprehensive threats assessments on communi-

ty, river, watershed, and faunal province spatial scales.

4. Predict how species and communities will change in

response to threats.

5. Work with the states and the U.S. Environmental Protection

Agency (USEPA) to modify water quality criteria, develop

new standards, and modify total maximum daily loads to

protect mollusk populations.

6. Advocate for consistent and effective enforcement of

environmental protection laws and regulations and evaluate

whether existing regulations adequately protect mollusks

and their habitats.

7. Develop an early detection and rapid response system for

new aquatic invaders.

8. Develop and publish protocols to avoid the spread of

disease and invasive species.

9. Describe the ecological and economic impact of mollusks

4 FMCS 2016

as vectors of parasites on fish, wildlife, livestock, and

human health.

Overview

Due to their varied life histories, sedentary nature, and

relatively poor dispersal mechanisms, mollusk populations are

susceptible to numerous biotic and abiotic stressors. For

example, understanding how contaminants affect mollusk

populations is complicated by residual contamination from

pollution and the lack of data on synergistic effects of multiple

contaminants on mollusks. Thus, even though an initial

stressor may be gone, other stressors may continue to

adversely affect mollusks and their habitats.

The tissues and shells of mollusks provide a long-term

record of past environmental conditions. For example, tissues

bioaccumulate contaminants (Naimo 1995; Coogan and La

Point 2008) and shells have been used to identify a suite of

past environmental events, such as climate and water quality

(Dunca et al. 2005; Rypel et al. 2008). Stressors on mollusks

are continually changing and research will be required to

identify and address the effects of new and existing stressors.

Examples of new stressors fall into categories of biotic

introductions (e.g., invasive snails, fishes) and abiotic

modifications (e.g., climate change, emerging contaminants)

of aquatic ecosystems. Since mollusks play an important role

in nutrient cycling and structuring food webs in aquatic

ecosystems (Atkinson et al. 2013, 2014a), clarification and

quantification of how mollusks respond to these stressors may

help develop criteria for healthy aquatic systems.

Success stories in emerging stressors

Because stressors are ubiquitous in aquatic systems, there

have been many opportunities to study their effects on mollusk

populations. In the 1990s and 2000s, considerable research

was completed on the effects of dreissenid mussels on native

mussels (e.g., Nalepa and Schloesser 2014). This has resulted

in several positive outcomes; for example, scientists now have

new tools to predict and perhaps prevent the spread of

dreissenids into new areas (Nalepa and Schloesser 2014;

Karatayev et al. 2015). These studies have also identified

refugia for mollusks in several systems where populations are

surviving despite the presence of dreissenids (Nichols and

Wilcox 1997; Crail et al. 2011; Zanatta et al. 2015). To our

knowledge, little research has been done on the effects of

dreissenids on snails. However, research has documented the

effects of invasive snails on native snails (e.g., Brown et al.

2008). We are unaware of research on the effects of invasive

snails on mussels.

Considerable progress has been made in understanding the

effects of abiotic stressors on mollusk populations. First,

mussels are now recognized as the most sensitive organisms

ever tested to the effects of ammonia (Augspurger et al. 2007;

Newton and Bartsch 2007; Wang et al. 2007a, 2007b). This

research prompted a revision of water quality criteria for

ammonia to be protective of mollusks (USEPA 2013). Second,

the impoundment of rivers with dams has often been cited as a

primary reason why many mollusk populations are fragmented

(Downing et al. 2010). Dams and channelization are under-

stood to be a leading cause for the extinction of at least 12

species of mussels and 45 species of snails (Haag 2009;

Johnson et al. 2013). Consequently, there is substantial interest

in removing dams to increase connectivity and improve

mollusk populations (Sethi et al. 2004; Nijhuis 2009; Haag

and Williams 2014). However, reproducing populations of

mussels have been documented downstream of barriers,

suggesting that river modifications should be considered on

case-by-case basis because dams may have a protective role for

some populations (Gangloff et al. 2011). Third, due to certain

life history traits (e.g., patchy distribution, limited mobility,

larval dependence on hosts, range fragmentation), mussels may

be especially susceptible to the effects of climate change. Many

of these life history traits may also adversely affect snail

populations (Johnson et al. 2013). In fact, research has shown

that many species of mussels are already living close to their

upper thermal limits and that there are physiological and

behavioral effects of climate change on mussels (Pandolfo et al.

2010; Archambault et al. 2013; Ganser et al. 2013). Advances

in modeling suggest dire consequences for mussel populations

as a result of climate change due to co-extirpation of mussels

and their hosts (Spooner et al. 2011).

Research to aid conservation and management

To assess and monitor the threats of stressors on mollusk

populations, data on tolerance limits (e.g., temperature, dissolved

oxygen, conductivity), sublethal effects, and habitat delineation

are urgently needed (e.g., International Union for Conservation

of Nature threats classification scheme). In addition, there are

limited data on the synergistic effects of stressors on mollusks

and on sensitivity of different taxa. Because the responses of

mollusks to stressors may vary by watershed, climate, habitat

type, taxa, and physiological state of mollusks, future studies

should evaluate the synergistic effects of multiple stressors across

species, life stages, spatial scales, and over varied environmental

conditions. Mollusks require an abundant supply of clean water

for physiological processes. Unfortunately, the quality and

quantity of water in many mollusk habitats is in jeopardy (Poff

et al. 1997; Arthington et al. 2006). Research is needed to

understand how demands on water supply will affect mollusk

communities and to develop instream flow criteria. We have

evidence that low flow periods—due to drought conditions in

Southern rivers—reduced species richness and relative abun-

dance of mussels (Haag and Warren 2008; Galbraith et al. 2010).

Scientists need to educate local water management districts on

how their practices (e.g., water extraction, diversions) may

adversely affect mollusk assemblages. Without this collabora-

tion, conflict between stakeholders can occur (Buck et al. 2012).

Endemic species (e.g., those limited to certain river reaches or

springs) are at risk of extirpation with even one major climate

5NATIONAL STRATEGY FOR FRESHWATER MOLLUSKS

event. Understanding the thermal limits of mollusks (and its

interactions with flow) across species and life stages will allow us

to forecast potential changes in mollusk assemblages due to

climate change. There also is a need to understand the direct

physical effects that biologists may have on mollusk assemblages

and the potential for introducing pathogens into established

populations.

Opportunities for conservation and management

Advances in new technologies can help researchers and

managers understand the effects of stressors on mollusk

populations. For example, environmental DNA (eDNA) is

currently being used to track the spread of invasive species

and could be used to detect mollusk populations in the future

(Olson et al. 2012; Pilliod et al. 2013). In a 5th order river,

New Zealand mudsnails (Potamopyrgus antipodarum) were

successfully detected using eDNA at densities as low as 11

individuals m�2 (Goldberg et al. 2013). This technology

could also be used to identify locations of at-risk populations,

allowing mangers to focus their conservation efforts. In

addition, the introduction of passive integrated transponder

(PIT) tags has allowed researchers to non-invasively monitor

the growth, survival, and movement of individual mussels

(Kurth et al. 2007; Newton et al. 2015). These techniques

allow scientists to identify and develop sensitive response

metrics from exposure to environmental stressors. Finally,

advances in testing methods and analytical techniques should

result in improvements across a range of state and national

water quality standards that could protect mollusk popula-

tions, similar to improvements made for ammonia.

Given that mollusks influence the ecology and economy of

fish, wildlife, and ultimately human health, scientists need to

provide managers with the tools (e.g., protocols for mollusk

management) to make decisions about how future and current

stressors and their interactions might affect the conservation of

mollusks. Efforts should also be made to support research on

the long-term effects of stressors because many are likely

surrogates for other emerging stressors.

Issue 3 – Understand and conserve the quantity and qualityof suitable habitat for mollusks over time.

Goal: Increase understanding of physical, chemical, and

biological characteristics of habitat to support sustainable

assemblages of mollusks.

Strategies:

1. Define habitat requirements at multiple spatial scales (e.g.,

organismal, population, community, river, watershed,

faunal province).

2. Define habitat requirements at multiple temporal scales

(e.g., seasonal, annual, long-term), including the quality,

quantity, and timing of ecological flows.

3. Quantify the amount of occupied and unoccupied habitat.

4. Identify and conserve habitats that will be resilient to

changing climates.

5. Reduce habitat fragmentation and increase connectivity of

historical habitats.

6. Conserve and restore habitat through land protection

actions such as easements, acquisitions, and landowner

agreements along riparian and upland areas within

watersheds.

7. Identify best habitat management and restoration practices

through evaluation, monitoring, and modeling.

8. Develop and test effective mitigation alternatives for

activities that affect habitat.

Overview

Habitat degradation and loss is a common threat to

mollusks, and often a major concern for sustainability of at-

risk mollusk populations (Lydeard et al. 2004; Strayer et al.

2004). Widespread construction of dams has resulted in vast

changes to habitats, flow and temperature regimes, and

ecological functions (Poff et al. 2007). Development of

restoration and mitigation tools for mollusk habitat, therefore,

is critical. For example, habitat loss is often listed as the most

significant threat in recovery plans for endangered mollusks,

but few plans provide guidance on habitat restoration and

instead focus on the nonetheless important protection of

diminishing and rare inhabited areas (e.g., USFWS 2004).

Compared to other charismatic or commercially-important

species (e.g., mammals, gamefish), the state of knowledge

needed to address conservation and management of habitats

for mollusks is poorly developed; yet this knowledge could

help explain other aspects of the ecosystem. Management of

mollusk habitat is further challenged by the complexity of

variables involved, regulatory constraints, and by multiple and

increasingly severe stressors on freshwater ecosystems (see

Issue 2) that vary across and within ecoregions.

Success stories in mollusk habitat

Although progress has been made toward understanding

habitat requirements of some mollusks—particularly mus-

sels—habitat needs of most species and life stages are not well

understood and remain a critical bottleneck for conservation

efforts (Johnson et al. 2013; Haag and Williams 2014).

Assessment and quantification of snail habitat has been

limited, but advances have been made toward understanding

habitat requirements in several aquatic systems (e.g., Evans

and Ray 2010; Calabro et al. 2013; Johnson et al. 2013). Such

understanding has enabled conservation gains for some

species. For example, the range of the endangered Tulotomamagnifica, a snail in the Coosa River, Alabama, had been

reduced by 99% (Hershler et al. 1990). Removal of dams to

restore riffle habitat, coupled with improved water releases at

hydroelectric dams and habitat protection, resulted in recovery

to 10% of its historical range and subsequent reclassification

from endangered to threatened (USFWS 2011).

6 FMCS 2016

For mussels, past research using traditional habitat

descriptors (e.g., velocity, particle size) was largely unsuc-

cessful at predicting occurrence or density (Strayer and Ralley

1993; Brim Box et al. 2002). More recent studies across a

variety of systems have suggested that certain complex

hydraulic variables (e.g., shear stress, Reynolds number) are

more predictive (Gangloff and Feminella 2007; Steuer et al.

2008; Zigler et al. 2008). While this has been a significant

accomplishment in quantifying physical habitat, mollusks

require more than physical habitat. Suitable patches of quality

habitat also require availability of mussel hosts, limited

predation, suitable water quality, and adequate food resources

(Newton et al. 2008).

Research to aid conservation and management

Future strategies to conserve or enhance habitat for

mollusks should focus on two areas. First, understanding

temporal and spatial patterns in mollusk habitat across

multiple scales and life stages will be critical in developing

effective conservation strategies. Predictive models that

quantify mollusk habitat and clarify functional habitat

attributes limiting mollusks are urgently needed. However,

development of habitat models is often challenged by a lack of

biological and environmental data, but recent advances in

technology (e.g., remote sensing, low cost hydraulic models)

and techniques for statistical and geospatial modeling of

habitat will facilitate future efforts. Depending on the factors

limiting mollusk habitat, these models may need to be scale-

specific (e.g., site, river, watershed, faunal province). Models

are also needed to understand the effects of dams, flow

alteration, and other anthropogenic stressors on habitat and to

forecast likely outcomes under differing management scenar-

ios and emerging issues such as competition for water

resources and climate change (Spooner et al. 2011). Additional

research is needed to describe the importance of spatial

arrangement and connectivity of quality inhabited and

uninhabited habitat patches to persistence of mollusk assem-

blages.

Second, research is needed to develop tools for imple-

menting and assessing conservation and restoration projects

which increase the amount of quality habitat for mollusks.

Partners may lack expertise, and often face difficult decisions

regarding efficient allocation of limited resources. Manage-

ment efforts such as artificial propagation may be futile if the

habitat is unsuitable due to other factors such as degraded

watersheds, and altered hydrology or hydraulic conditions.

Evaluation and monitoring to determine the effectiveness of

management actions to protect or mitigate lost or impaired

habitats is essential for improving methods in an adaptive

management framework. Improvements in methods for

restoration or mitigation are likely to be incremental and

require long-term commitments to fully understand ecological

outcomes.

Opportunities for conservation and management

Conservation of mollusks depends greatly on continued

advances in conservation of their habitats. Opportunities for

improving mollusk habitat will range from local to watershed-

level efforts. In some systems such as the Upper Mississippi

River, scientists, managers, and engineers are in the planning

stages of constructing habitat restoration projects to benefit

mussels by altering local physical and hydraulic conditions.

Such projects present substantial opportunities for understand-

ing the important features of mussel habitat and developing

restoration and mitigation tools. Larger scale projects that

might target mollusk habitat include improving land-use in the

watershed (e.g., controlling sediment and contaminant inputs)

and restoring a more natural hydrograph and ecological flows

in rivers. Targeted dam removal, practices that improve water

and sediment quality, and repairing altered or channelized

streams are other actions that may improve mollusk habitat.

Habitat restoration projects for many drainages are underway

through partnerships such as the National Fish Habitat Action

Plan and National Fish Passage programs of the USFWS and

state and non-governmental initiatives. Biologists and manag-

ers should engage and partner with these organizations to

include objectives for restoring mollusk habitat. Because

mollusks can provide beneficial ecological services (see Issue

6), and can function as keystone species (Geist 2010), there

can be substantial synergy between efforts to restore habitat

and conservation goals for other ecosystem components.

Issue 4 – Understand the ecology of mollusks at theindividual, population, and community levels.

Goal: Increase fundamental knowledge of the biology of

mollusks so managers can more effectively conserve them.

Strategies:

1. Describe life history and host-species relationships at the

appropriate scale.

2. Define environmental and nutritional requirements neces-

sary for physiological maintenance, reproduction, and

persistence of all life stages.

3. Describe the ecological functions of mollusks in the

environment.

4. Increase knowledge of negative and positive interactions

among mollusk species.

5. Increase understanding of demographic, genetic, and

physiological characteristics that influence long-term

population viability.

6. Encourage development of population viability analyses to

better predict species’ persistence.

7. Develop population goals for managing rare and common

species.

Overview

Although we have learned a great deal about the ecology of

mollusks since the 1998 National Strategy, we still have much

7NATIONAL STRATEGY FOR FRESHWATER MOLLUSKS

to learn. Most work has concentrated on mussels, but we need

more information on their demography, host-use and move-

ment, nutritional needs, habitat requirements for juveniles,

specific water quality limits, and sensitivity of all life stages to

multiple stressors. Snails have received considerably less

attention and there are significant data gaps in many areas

including life history, population demography, habitat require-

ments, species interactions, and ecological function. Under-

standing the ecology of mollusks at the individual, population,

and community levels is needed to design effective conserva-

tion and restoration strategies.

Success stories in mollusk ecology

Summaries of the ecology of mussels are contained in two

books (Strayer 2008; Haag 2012) and several review articles

(Vaughn and Hakenkamp 2001; Vaughn et al. 2008; Haag and

Williams 2014). Studies have begun to reveal the unique traits

that mollusks possess that enable them to cope with variable

environments. For example, research suggests that mussels can

change their feeding strategy in response to varied food

sources by feeding across trophic levels (e.g., bacteria vs.

phytoplankton) or mechanistically (e.g., pedal vs. filter

feeding) (Vaughn et al. 2008; Newton et al. 2013). Mussels

also have a variety of responses to cope with environmental

contaminants, such as varied detoxification capabilities

(Newton and Cope 2007). Improved understanding of the

strengths and weaknesses of survey methods has expanded our

understanding of populations and population processes

(Strayer and Smith 2003). For example, increased use of

excavation techniques that enhance detection of smaller

mussels can improve demographic information. Demographic

models have been developed for some rare species (Crabtree

and Smith 2009; Jones et al. 2012) leading to better

understanding of their ecology. Use of conservation genetics

can help determine long-term viability of populations (Jones et

al. 2006). We have also developed temperature criteria for

many species and life stages of mussels (Spooner and Vaughn

2008; Pandolfo et al. 2010, 2012).

Research to aid conservation and management

We have made substantial progress in understanding mussel

life histories and host-species relationships over the past two

decades (e.g., Barnhart et al. 2008). However, there are still

substantial data gaps. We have host-use data for only about a

third of the North American mussel fauna, and much of that

information is incomplete (Haag and Williams 2014). Repli-

cated, robust studies that document the breadth of host-use

across co-occurring fishes are needed (Haag and Williams

2014). Once potential hosts are known, there is also the

challenge of determining which hosts are most important in

natural systems. We also need to understand patterns in juvenile

mussel dispersal that host movements provide. Studies using

PIT tags and other novel techniques will increase knowledge in

this area. We also need data on sperm dispersal in the field and

reproductive success at low mussel densities, which may be a

bottleneck for the conservation of small and isolated popula-

tions. In snails, representative pleurocerid and pulmonate

species have been the subject of life history studies (e.g.,

Huryn et al. 1994; reviewed in Brown et al. 2008), but we need

more information on reproductive biology of most species.

While vast progress has been made understanding habitat

requirements of mussels in the last decade (see Issue 3), more

work is required. In particular, we have limited knowledge of

the habitat needs of juveniles. We also need to examine

sensitivities to habitat perturbation in terms of multiple

stressors, rather than single stressors (Shea et al. 2013). There

is evidence that the early life stages of mussels are particularly

sensitive to contaminants, such as un-ionized ammonia and road

salt (Augspurger et al. 2003; Newton et al. 2003; Gillis 2011),

but there are thousands of contaminants that have not been

evaluated (see Issue 2). Sensitivities of mussels to the effects of

climate change are just becoming apparent, but there is much

more to learn (Ganser et al. 2013; Archambault et al. 2014). We

also have inadequate knowledge of nutritional requirements of

mollusks across life stages (Haag 2012).

Because we cannot determine the ecological requirements

and sensitivities for all mollusk species, one approach is to

look at requirements and tolerances of guilds or suites of

species with similar traits (Statzner and Beche 2010; Lange et

al. 2014). For example, groups of mussel species with similar

thermal traits respond differently to drought-driven thermal

stress (Atkinson et al. 2014b). Thermal and life history traits of

species guilds have been combined to make flow recommen-

dations for mussels, based on the traits and criteria for the most

sensitive species (Gates et al. 2015). Comparative life history

studies across populations should be performed on represen-

tatives from the major snail families to see if ecological

requirements and sensitivities are similar across species or

genera (e.g., Huryn et al. 1994).

Mussels are largely filter-feeding omnivores that feed

across trophic levels on bacteria, algae, and other suspended

material (Vaughn et al. 2008). Many mussels live in multi-

species aggregations (i.e., mussel beds); thus, species

interactions should be important in their communities. Most

studies have concentrated on negative species interactions

(competition) with varied results (Haag 2012), and more work

on interactions between and within mussel species is needed.

In particular, studies are needed on when and where food and

space may be limiting. In addition, positive (facilitative)

interactions may also be important in mussel communities,

similar to marine bivalves and other groups of freshwater filter

feeders (Cardinale et al. 2002; Spooner and Vaughn 2009;

Angelini et al. 2011). As with the ecological requirements, the

largest data gap is for early life stages. For example, juvenile

survival rates and the effect of critical factors such as

parasitism and predation rates are mostly unknown. Research

is also needed to answer questions about the ecological

interactions of adult and juvenile mussels (e.g., Do adults

facilitate juvenile survival by providing a protective environ-

8 FMCS 2016

ment? Does the community of adults and different species

create micro- or macro-environments of suitable habitat for

juveniles? Do adults compete with juveniles for limited food

resources, or do adults provide organic substrates that help

build food resources for juveniles?).

Similar to mussels, research on snails is needed to better

understand species and ecosystem interactions. Snails can be

the most abundant invertebrate grazers in streams in the

Southeastern U.S. with large ecosystem effects (Richardson et

al. 1988; Hill 1992). Snails can reduce periphyton biomass and

alter algal community composition, which can lead to changes

in primary production (Brown et al. 2008). For example, the

invasive New Zealand mud snail consumes up to 93% of

primary production in streams (Hall et al. 2003), and the

invasive Chinese mystery snail, Cipangopaludina chinensismalleata, can influence algal community structure (Johnson et

al. 2009). Aside from a few representative pleurocerids, much

of the work on species interactions in snails has been on

interactions with invasive snail species. For example, the

Chinese mystery snail affects native snails by reducing their

populations, modifying their habitats, and increasing parasite

pressure (Harried et al. 2015).

We know little about how to measure the relative health of

mollusk populations, or what constitutes a self-sustaining

population. Traditional measures of mussel populations (e.g.,

density, species richness) may not be sensitive enough due to

long response times and life spans (Newton et al. 2008). Lack of

demographic data at the species and community levels

complicates resource management. Few studies describe

demographics (e.g., recruitment, age structure), species richness,

and species evenness—metrics that might be used to evaluate

mussel community health (but see Villella et al. 2004; Haag and

Warren 2010; Newton et al. 2011). Long-term monitoring

studies that describe how demographics vary over space and

time are also lacking. Multi-metric indices have been developed

for sensitive fish and invertebrate communities to assess the

health of these communities in rivers and lakes. However,

investigations into the sensitivity of mollusks to contaminants

and habitat perturbations are relatively recent. Experimental

multi-metric indices have been developed for mussel commu-

nities in several states and regions, but have not been rigorously

tested. Unfortunately, we know little about the physiological or

genetic characteristics that promote long-term population

viability (but see Gough et al. 2012; Archambault et al. 2013;

Gray and Kreeger 2014). The lack of these data limits our

ability to assess how human activities might adversely affect

mollusk populations.

Opportunities for conservation and management

Robust data on mussel abundance, distribution, and status

are starting to be obtained in many drainages, providing

critical data for conservation efforts. Meta-analyses of these

data within and across drainages should uncover interesting

demographics and inform management strategies. However,

life history data for many imperiled species (e.g., Hydrobiidae)

are lacking and should be addressed. Techniques for

documenting host-use by mussels are expanding, including

the use of advanced microscopy and genetics. Advances in

analytical chemistry now make it possible to measure

environmental contaminants that are biologically active at

low concentrations (e.g., pharmaceutical products, nanoparti-

cles, hormones). Coupled with an increased awareness of

environmental contaminants in waterbodies, there are many

opportunities to understand the effects of contaminants on

mollusks and their hosts. The success of propagation efforts

has made multiple species and life stages available, greatly

expanding the potential for toxicological testing with mol-

lusks. Advances in population and environmental modeling

tools provides an opportunity to understand and predict the

effects of multiple, interacting stressors on mollusk popula-

tions as well as developing effective and cost-efficient

management strategies.

Issue 5 – Restore abundant and diverse mollusk populationsuntil they are self-sustaining.

Goal: Conserve and restore viable populations and

communities of mollusks.

Strategies:

1. Develop population and community indices to monitor and

evaluate sustainability over time.

2. Develop conservation and restoration plans (e.g., reintro-

duction or augmentation) at the river, watershed, and faunal

province levels.

3. Implement restoration that results in self-sustaining popu-

lations and does not adversely affect resident fish and

mollusk populations and their habitats.

4. Maintain a database of translocation, propagation, and

stocking events.

5. Identify uniform methods and metrics for monitoring

outcomes of augmentations and reintroductions.

Overview

Mollusks have a 74% imperilment rate, with many species

that rank among the most imperiled animals on the continent

(Johnson et al. 2013; J.D. Williams, Florida Museum of

Natural History, personal communication). In addition to

species considered imperiled and federally endangered, many

wide-ranging and common species also are no longer

prevalent. This loss diminishes biodiversity and affects the

ecosystem services mollusks provide in functional aquatic

systems (Spooner and Vaughn 2008). Moreover, diverse and

abundant mollusk communities are more robust and resistant

to invasion by non-native species (Strayer and Malcolm 2007).

High mussel species diversity is found in communities with

rare species and good water quality, so community data would

be required for understanding the sustainability of all

populations. Population trend analyses would also give

9NATIONAL STRATEGY FOR FRESHWATER MOLLUSKS

biologists essential data to understand the ecological causes of

these declines.

To increase the abundance and distribution of mussel

populations, more needs to be known about demographics,

viability, and genetics of fragmented populations (Haag and

Williams 2014). Indeed, the role of population fragmentation

on species and community declines must be addressed.

Mollusk restoration efforts have largely focused at the site

scale for individual species; rarely have these efforts been

addressed at the population, much less community, level. It is

imperative that defined population goals are established and

that methods are developed to monitor populations and

communities over time, given the pervasiveness of disjunct

and isolated populations with limited recruitment. Conserva-

tion strategies should maximize the benefits of disjunct

populations to the overall species conservation (i.e., tracking

genetic diversity of animals used for propagation, mixing of

animals from adjacent shoals). This includes establishing

accessible databases to hold unpublished community data.

Success stories in mollusk restoration

With the passage of the ESA in 1973 and the current listing of

91 mussel and 31 snail species in North America, research

facilities began to investigate the life histories of individual

species. The goal of these investigations was the development of

propagation techniques for restoring imperiled species through

stocking of cultured progeny. Since the late 1990’s, several

propagation programs throughout the U.S. have been successful

in propagating and stocking mollusks. Research efforts for

mussels have identified suitable diets, feeding regimes for adult

broodstock and juveniles, host-species, and numerous culture

and grow-out systems (e.g., Barnhart 2006; Gatenby et al. 2013;

Mair 2013). These efforts were often associated with stocking

programs aimed at restoring mussel beds adversely affected by

damages from chemical spills or other instream activities such as

bridge replacements (Morrison 2012; Morrison et al. 2013; Lane

et al. 2014). Propagation technology has improved greatly over

the past 20 years and some mussel species have been propagated

at a production scale (Neves 2004; Haag and Williams 2014).

Over a dozen federal and state culture facilities now exist in the

Midwest, mid-Atlantic, and Southeast regions of the U.S.

Examples of culture and population restoration efforts include

several imperiled mussel species in the Tennessee River

drainage, the Upper Mississippi River drainage, and several

drainages in Missouri (Barnhart 2002; Davis 2005; Carey et al.

2015). Culture facilities and population restoration programs are,

however, absent from other parts of North America, and some

highly imperiled taxa are virtually unstudied (e.g., the endemic

springsnails, Order Rissooidea). Evidence of self-sustaining

populations of mussels from propagation efforts is generally

lacking due to their longevity and slow population response

times. Propagation of snails is fairly recent, with only a few

programs available. For instance, Alabama has been culturing

several species of pleurocerid snails and is conducting population

restoration activities in several streams (P.D. Johnson, Alabama

Aquatic Biodiversity Center, personal communication).

Research to aid conservation and management

The focus of most mollusk restoration programs has been

on recovery of endangered species, primarily due to limited

funding. Managers must strive to obtain the resources

necessary for community restoration. Community restora-

tion—including rare and common species alike—will restore

the broad range of ecosystem services mussels contribute to

aquatic systems (Spooner and Vaughn 2008; Vaughn et al.

2008). Further, restoration of healthy populations increases the

resiliency of populations and enhances the probability for

successful restoration efforts (Sethi et al. 2004; Zanatta et al.

2015). Learning how to propagate mollusks across taxonomic

groups will allow managers to increase abundance and

distribution of populations, contributing directly to recovery

of imperiled species which are usually found with common

species. Coupled with efforts towards defragmenting habitats

and populations, the status of some imperiled species may

improve to the point of precluding their listing under the ESA

while enhancing the probability of recovering federally listed

species.

Stressors that affect entire assemblages of mollusks are

often habitat alteration-based, and include population frag-

mentation, non-point and point source runoff, environmental

(e.g., chemical spills, extreme droughts) and demographic

(e.g., altered fertilization and recruitment rates) stochasticity,

and altered flow regimes (Lande et al. 2003; Haag and

Williams 2014). Contaminants may affect mollusks differently

depending on species, life stage, and habitat conditions. Since

mollusks appear to be some of the most sensitive of all aquatic

organisms to some stressors, entire communities are vulner-

able (see Issue 2). The effects of physical habitat changes on

mollusk communities are needed at a scale suitable for

management (see Issue 3). Maintenance of adequate flow

regimes is important for managing mollusk populations. There

are also numerous unexplained community-level ‘‘enigmatic

declines’’ in mollusk populations that need to be addressed

(Haag 2012).

Many aquatic ecosystems rely on the index of biotic

integrity (IBI) for understanding biotic and abiotic interac-

tions. Mollusk data are often lacking in IBIs or at best, are

included only at the taxonomic class level (e.g., Calabro et al.

2013). Incorporating mollusk biodiversity into IBIs and/or

developing new IBIs would create stronger predictions of

aquatic communities and foster tracking the status of

mollusks over time. Mollusk-specific IBIs might be devel-

oped to provide a quantitative means of evaluating the

relative health or value of mollusk assemblages, as a tool for

conserving mollusk resources including measuring impacts,

assessing the efficacy of restoration techniques, and inform-

ing regulatory tasks. For example, scientists in the Upper

Mississippi River have created a mussel community assess-

10 FMCS 2016

ment tool to explore metrics to assess mussel community

health (Dunn et al. 2012).

Basic propagation technology (e.g., culturing, feeding,

growing out) and protocols for determining genetics, handling,

and quarantining mussels are generally available, but may

need to be revised to meet species-specific needs (Gatenby et

al. 2000; Cope et al. 2003; Jones et al. 2006). However, there

is a need to improve grow-out technology, in addition to

identifying the best host species and understanding the

physiological requirements to improve growth and survival.

Most mussel propagation research has been conducted on

Lampsilines, but efforts need to be expanded to include

Anodontines and Amblemines, which include many imperiled

species. Snail propagation research is still in the early stages of

development, partly due to a paucity of ecological and basic

life history data (Johnson et al. 2013). The diversity and

environmental sensitivity of some snails (e.g., Rissooidea,

Pleuroceridae) also pose challenges for developing propaga-

tion methods. In addition, obtaining broodstock is often a

limiting factor in culture efforts. Research to facilitate natural

reproduction in the laboratory would allow culturists to

circumvent the need for wild broodstock. In vitro culture of

mollusks could reduce labor and space costs by eliminating the

need for mussel host species and may reduce the risk of

disease in hatchery or culture settings.

Minimizing risks to aquatic systems from population

reintroduction efforts must become a priority (Olden et al.

2010). We should aim to reduce risks to resident populations

of mollusks and host-species and carefully consider genetic

diversity of both resident and stocked mollusks. Choosing

taxonomically well-studied populations for activities (Jones et

al. 2006; Hoftyzer et al. 2008) should lessen the threat from

inbreeding and outbreeding depression. Restored populations

should be closely monitored to detect any potential negative

interactions that may occur. Because restoration of natural

populations is a primary management goal, it is important to

identify and publish metrics for monitoring outcomes of

management actions to promote development of uniform,

repeatable, and successful methods.

Opportunities for managing and restoring populations andcommunities of mollusks

Coupled with further advances in the biology and ecology

of mollusks, malacologists are calling for comprehensive and

universal protocols by resource agencies engaged in restoring

mollusks through stocking programs (Jones et al. 2006;

Hoftyzer et al. 2008; McMurray 2015). For example, the

development of a comprehensive course on propagation of

mussels—held at the National Conservation Training Center

(NCTC) of the USFWS—has trained biologists to develop

propagation programs for mussels. We have a few examples of

restoration aimed at the community-level; however, more

restoration efforts should target mollusk communities. Indeed,

several recovery plans, strategies, and restoration plans

approach restoration through methodical and coordinated

planning across geographic scales (e.g., Hartfield 2003;

USFWS 2004, 2010, 2014; Cumberlandian Region Mollusk

Restoration Committee [CRMRC] 2010). These plans prior-

itize species, streams, and activities for conservation and

management efforts, which will aid managers in restoring

populations and communities at the watershed level. Restora-

tion plans typically stress reintroducing extirpated populations

over augmenting extant ones for two reasons: 1) recovery can

only be achieved by creating additional populations and 2)

augmentations have unique inherent risks to existing popula-

tions (CRMRC 2010; Haag and Williams 2014). Stream

reaches and lakes affected by legacy environmental or human

development issues can be restored to promote connectivity of

mollusk populations (Newton et al. 2008).

In certain cases, federal listing under the ESA (and possibly

even state listing) has been construed as a hindrance to

recovery. This is despite the substantial benefits that Section 6

of the ESA and the State Wildlife Grants Program (SWG) have

had on state funding for recovery actions for federally listed

mollusks and the high profile that federal and state listing has

provided. Regulatory burdens may be increased due to

construction projects with a federal connection (under Section

7 of the ESA) or recovery activity implementation may have

time constraints imposed due to permitting issues (Section 10).

These issues can be ameliorated with better communication and

relationships among USFWS, state, and other partners. The

Section 7 process can and should be implemented to further

recovery of species potentially affected by construction projects.

Discussions with state biologists on how to implement recovery

activities while reducing regulatory requirements will serve to

expedite recovery of listed species.

Issue 6 – Identify the ecosystem services provided bymollusks and their habitats.

Goal: Improve science-based consideration of the social

and economic values of mollusk communities and functioning

aquatic systems.

Strategies:

1. Describe ecosystem services provided by mollusks to

humans and river ecosystems.

2. Develop and publish the social and economic values of

healthy mollusk communities to society.

3. Update the values and replacement costs of mollusk

communities.

4. Publish a comparison of mollusk replacement costs with

other biologically engineered mitigation alternatives.

Overview

Ecosystem services are the benefits that humans derive from

healthy ecosystems. These include provisioning services obtained

directly from the ecosystem such as water, food, and timber;

regulating services such as water purification, climate control,

carbon storage, and pollination; and cultural services which are

11NATIONAL STRATEGY FOR FRESHWATER MOLLUSKS

the benefits that people obtain through tourism and recreation,

aesthetic experiences, or spiritual enrichment. Biologically

complex freshwater systems provide many important ecosystem

services that benefit society such as provisioning of clean water

(water filtration and nutrient processing), recreation, and

ecotourism (Brauman et al. 2007; Dodds et al. 2013). Mollusks

provide all of these services for humans ranging from food, water

filtration and nutrient processing, to use in a multi-billion dollar

pearl jewelry industry. Mussels were also culturally important to

Native Americans as a food source and for use in jewelry, pottery,

and tools (Klippel and Morey 1986; Haag 2012). Harvest of

mussels is a reserved treaty right in the U.S. for some Native

American tribes (Brim Box et al. 2006). Thus, ecosystem services

provided by mollusks are important for human wellbeing.

Success stories in ecosystem services provided by mollusks

Researchers have learned a great deal about the services that

mollusks provide to lakes and rivers over the past 20 years.

Snails are important grazers and detritivores that support

decomposition, influence algal and bacterial communities, and

are an important food source for many fish, reptiles, and birds

(Hall et al. 2006; Brown et al. 2008). Mussel shells provide

habitat and refugia for other organisms, including benthic algae,

macroinvertebrates, nesting fish, and other mussel species

(Vaughn 2010). Mussels can aerate sediments and alter sediment

stabilty and erosional processes, further improving habitat for

mussels and other organisms (Zimmerman and de Szalay 2007;

Allen and Vaughn 2011). Assemblages of common mussel

species also improve conditions for rare mussel species that are

typically found within larger assemblages (Spooner and Vaughn

2009). Filtering mussels remove seston from the water column,

which can decrease water treatment costs and improve water

quality (Newton et al. 2011). Mussel beds create biogeochemical

hotspots via nutrient excretion and storage (Strayer 2014;

Atkinson and Vaughn 2015). Where nutrients are limiting,

fertilization by mussel excreta supports the rest of the food web,

leading to increases in benthic algae, macroinvertebrates, fish,

and even riparian spiders (Allen et al. 2012; Atkinson et al.

2014c). Mussel-provided nutrients can also alter algal compo-

sition, leading to decreasing blue-green algae populations and

increasing water quality (Atkinson et al. 2013). There are few

comparable data on ecosystem services provided by snails.

Research to aid conservation and management

Although we have a better understanding of the ecological

services that mollusks provide to aquatic systems, we now

need to quantify how losing these services may affect aquatic

systems, and the resulting economic and social consequences

to humans. A good place to start would be to partner with

scientists and managers who have conducted natural resource

damage and restoration assessments for oyster beds (Beck et

al. 2011). Discussions with those who have performed rare

species valuations would be especially valuable (Richardson

and Loomis 2009). These assessments should explicitly

include the preferences and perceptions of major stakeholder

groups (Menzel and Teng 2010). Considering data on

economic values without also considering social values will

underestimate mollusk ecosystem services (Seppelt et al.

2011). Gathering and analyzing these data will require that

biologists collaborate with social scientists and economists

skilled at these analyses.

Opportunities for conservation and management

Nutrients stored in mussels (particularly in shells) are

retained in the system long-term because most mussels are

relatively long-lived. Thus, nutrients recycled by mussels are

incorporated into the stream food web rather than being

transported downstream (Atkinson et al. 2014c). This is also

likely the case for snails, but research is needed to quantify

their nutrient cycling and storage capabilities. Although

nutrients retained in this manner in one river may seem

insignificant, summed across multiple watersheds, this bio-

logical nutrient retention could help mitigate the effects of

nutrient pollution. Large-scale production of mussels could be

used to restore their biomass to enhance nutrient abatement. In

addition, outreach and communication about the value of

mollusks for providing important ecosysytem services should

be developed to increase public support for protecting their

populations (see Issue 7). Ongoing research in mollusk

physiology (see Issue 4) will help quantify their contribution

to improving water quality and reducing treatment costs.

Both marine bivalves and freshwater dreissenids can increase

nitrification and denitrification in sediments by biodepositing

organically rich feces and pseudofeces (undigested particles) on

which bacteria feed (Bruesewitz et al. 2009; Kellogg et al. 2013;

Smyth et al. 2013). Mollusks should also have strong effects on

coupled nitrification-denitrification by biodepositing organic

material, thus increasing rates of both processes, and by

bioturbating sediments as they move (Vaughn and Hakenkamp

2001). However, the effects of mollusks on nitrification-

denitrification need to be quantified.

Issue 7 –Strengthen advocacy and build support for theconservation of mollusks and their habitats.

Goal: Increase information sharing and communication

among citizens and decision-makers at multiple levels (e.g.,

local, state, regional, national, international) regarding con-

serving mollusk resources.

Strategies:

1. Develop a formal communication plan to guide conserva-

tion of mollusks into the future.

2. Develop science-based communication tools for local

decision makers to build organizational and public support

for land-use practices that support healthy aquatic systems

including mollusks.

12 FMCS 2016

3. Develop communication and outreach materials targeting

the general public, and to build awareness, appreciation,

and support for conservation of mollusk resources.

4. Empower citizens with necessary outreach materials to

advocate for consistent and effective enforcement of laws

and regulations or to develop new regulations.

5. Recruit communication specialists to the mollusk conser-

vation community.

6. Work with international partners to develop a global

strategy for the conservation of mollusks.

7. Increase collaboration with other aquatic societies.

8. Increase collaboration with other resource agencies and

resource groups.

Overview

Knowledge of the current conservation status of mollusks

is limited to experts and others that have been educated as a

result of acute and chronic impacts to the environment. To

conserve mollusks and their habitats, the risks and benefits

outlined in this manuscript need to be effectively communi-

cated to decision-makers, conservation groups, and the public.

There is a growing need for information that will aid public

advocacy for environmental legislation. Most of the commu-

nication by scientists is done formally through scientific

presentations of research, technical reports, and peer-reviewed

publications, or informally through face-to-face meetings,

email, and social media. While sharing scientific knowledge

and new discoveries is important, science writing is often

directed at a small and specialized audience. To reverse

declines in habitat quality, biodiversity, and budgets for

mollusk conservation, we need to work with others who can

affect change. Our communication skills and tools need to be

broadened to convey information that will reach other

conservation groups, the public, and policy makers.

Success stories in advocacy and communication

Partnerships and communication have greatly increased

support for mollusk conservation. Hundreds of papers, agency

reports, and government documents on mollusks have been

published (Figure 1). Publication and access to these data has

resulted in many of the success stories highlighted throughout

this manuscript. Advances in computer information transfer

and online access to publications, unpublished reports, and

other data have played a role in facilitating conservation and

management efforts. For example, numerous websites for

mollusks now exist that aid managers, researchers, and the

public in accessing information and providing timely updates

on imperiled species (Barnhart 2010; Dillon et al. 2013; Graf

and Cummings 2015; Watters and Byrne 2015). Global

partnerships can benefit this National Strategy through many

mechanisms including external pressures on policy, increased

knowledge, and new technologies.

In 2009, the FMCS organized a symposium with an

international focus that brought together researchers and

managers engaged in mollusk conservation. Several on-going

international collaborations resulted from this effort. Other

international meetings aimed at mollusk conservation have

occurred since 2009, strengthening communication and

collaboration among the international mollusk community.

Likewise, scientists working with mollusks have had a marked

presence at a variety of non-mollusk conferences (e.g.,

American Fisheries Society, Society for Freshwater Science,

American Society of Limnology and Oceanography), raising

awareness and advocacy for the conservation of mollusks and

their habitats. These examples show that partnering with other

natural resource or taxonomic groups can leverage resources

and expertise in protecting aquatic resources.

The mollusk conservation community has also developed

many outreach and communication tools to increase awareness

and advocacy of the global plight of mollusks. For example,

children’s books and informational posters have been created

on the biodiversity of mollusks, ecosystems, watersheds, and

the history of pearl culture. A large floor-model display of the

biology of mollusks has facilitated outreach at conferences and

videos on the biology of mussels are available (Barnhart

2010). The quarterly online FMCS newsletter, Ellipsaria(http://molluskconservation.org/Ellipsaria-archive.html), con-

tains outreach materials and is an important communication

tool for researchers, managers, and the public.

Communication to aid conservation and management

To have a greater effect on environmental and societal

issues that threaten the health and resiliency of aquatic

ecosystems, our passion for mollusks needs to reach those

outside the mollusk conservation community. The public

needs to understand how important mollusks are in creating

healthy environments for a variety of other animals, including

humans (see Issue 6). Recognizing that landowners often have

direct influence on the management of riparian zones, outreach

to farmers and urban planners should be prioritized. We should

also increase collaboration with human health and industry

groups to build partnerships to reduce environmental threats to

humans and aquatic resources.

In today’s ever-changing communication landscape, there

is considerable competition for news and information.

Compelling stories appropriate for varied media outlets are

needed to ignite passion for conserving our limited freshwater

resources. To ensure that information reaches the most

influential audiences, we must develop strategic and creative

ways to deliver our message. Specifically, a communication

plan should be developed to address topics such as (1) training

scientists and managers to effectively communicate with the

public; (2) recruiting communication specialists as partners in

creating effective communication that achieves conservation;

(3) developing presentations and outreach materials aimed at

public forums (e.g., video and film; outreach with zoos,

aquaria, and museums), policy makers (e.g., graphics showing

ecosystem services provided by mollusks, graphics highlight-

13NATIONAL STRATEGY FOR FRESHWATER MOLLUSKS

ing cost-benefit analyses of restoring mollusk assemblages),

and/or news media (e.g., blogs, news articles, press releases);

(4) developing a more visible presence on social media; and

(5) working with non-traditional groups outside the field of

conservation.

Opportunities for communication to conserve molluskassemblages

Partnerships offer opportunities to share resources and

increase awareness across groups and individuals that other-

wise would not be reached through the mollusk community

alone. There are many opportunities to partner with others in

the international community who are also working to reverse

declines in mollusks and restore habitats. Joint research

projects with international colleagues should result in enhanced

opportunities to restore mollusks through sharing of knowledge

and expertise. Collaboration on targeted communication

campaigns can strengthen public advocacy and political

support for conservation over the global landscape.

There are many ways to deliver a message, depending on

the content of the message and the audience. Since the impact

of major storms such as hurricanes Sandy and Katrina, there is

growing awareness and support by the public, the media, and

policymakers to restore landscapes that are resilient to rising

water levels and flooding. The mollusk community should

consider this enhanced awareness when promoting the

ecosystem services provided by mollusks (see Issue 6) and

to promote the benefits of intact, connected aquatic habitats

(Issue 3) for protecting human lives and property, as well as

the environment—facts that may be more compelling to

audiences than protecting rare mollusks.

Developing a communication plan which addresses the

issues outlined in this manuscript will make it easier to

communicate the value of protecting and restoring mollusks

and their habitats. There are many online examples of

communication plans; however, it may be cost effective to

enlist the help of professionals. In addition, science writing

and environmental journalism programs often seek opportu-

nities for students to work with scientists to communicate their

message. Sharing our conservation message with the public,

government representatives, the media, conservation groups,

landowners, and landscape planners must be a priority to

ensure conservation of mollusks is relevant to society.

Issue 8 – Educate and train the conservation community andfuture generations about the importance of mollusks toensure conservation efforts continue into the future.

Goal: Provide a suite of training opportunities to the

greater conservation community, and inspire future genera-

tions to work on the conservation of mollusks.

Strategies:

1. Develop and recommend a list of key skills and

competencies for mollusk conservation biologists and the

supporting disciplines such as communication.

2. Develop and recommend new coursework for the study and

conservation of mollusks based on the skills and compe-

tencies identified in Strategy 1.

3. Manage a database of training courses, internships, details,

and other professional opportunities for students and

practicing professionals to gain hands-on experience with

mollusk conservation.

4. Provide travel funds for FMCS members to attend training

courses, outreach events, educational institutions, college

fairs, and job fairs to encourage students interested in

biology and natural history to consider careers in mollusk

conservation.

5. Develop a FMCS grant program for students and other

conservation groups to advance the conservation of

mollusks through research and management activities.

Overview

Growing concern about the future of mollusk conservation

has elevated education and training to important issues. Many

long-time specialists in mollusk conservation are nearing

retirement and retention of their expertise in the scientific

community is needed for successful mollusk conservation.

Efforts to conserve mollusks could be undermined by this loss

of institutional knowledge and are further confounded by the

fact that documenting the long-term success or failure of

conservation actions may not be realized for decades or longer.

Many species are long-lived, slow growing, and have low

recruitment rates; knowledge gained from conservation efforts

will require long-term monitoring beyond the time-frame of

funded projects or careers.

Additionally, the science and technology underlying

conservation actions are changing rapidly. Thus, it is

imperative that we provide professional and academic training

opportunities for the conservation community to inspire the

next generation of scientists to pursue careers in mollusk

conservation. Even given their high degree of imperilment,

mollusks have been highly under-represented in publications

compared to vertebrates (Lydeard et al. 2004). We also

recognize that other areas of expertise and skills beyond the

sciences are needed to ensure mollusks persist and to help

conservationists deliver their message (see Issue 7).

Success stories in mollusk education

Currently, there is no group of individuals charged with

achieving these strategies. Rather, this issue largely overlaps

with the activities of FMCS committees (e.g., outreach,

symposium, information exchange, awards). The FMCS

actively sponsors annual educational events including sympo-

sia and workshops that provide opportunities to share the

current state of the science with the conservation community

and provide a venue for students to share their research.

Symposia and workshops have covered a range of themes,

14 FMCS 2016

including outreach, propagation, snail ecology, regional

mussel identification, environmental flows, climate change,

and dam removals. The FMCS offers travel awards to support

student attendance at symposia and provides monetary support

to regional mollusk conservation groups.

Formal courses are now being offered by federal agencies

on mussel conservation biology and propagation (e.g., NCTC).

Many states and private conservation organizations also offer

short courses, workshops, and training in mussel identification,

sampling techniques and protocols, and methods for handling

mollusks. A few states have developed guidelines, minimum

standards, and tests that certify scientists before they receive a

scientific collecting permit and conduct mollusk work within

their boundaries (e.g., Ohio, Pennsylvania, West Virginia).

Several universities offer graduate level classes and research

opportunities in mollusk conservation. However, training

opportunities for snail identification or biology remain

infrequent and informal.

Opportunities for education to enhance molluskconservation

The FMCS should coordinate and act as a clearinghouse

for relevant educational and training opportunities in mollusk

conservation. The FMCS could set up a grant program to

support conservation and communication projects by stu-

dents and conservation groups, and develop new workshops

that reflect the emerging science needed for effective

conservation.

Few universities offer undergraduate classes on mollusks

beyond a basic exposure in invertebrate zoology. There has

been a gradual shift from courses on organismal biology,

biodiversity, and taxonomy to courses on microbiology and

theory at academic institutions (e.g., Greene 2005). This

focus is needed to invigorate the study of mollusks and

capture students’ interest early in their educational career. In

addition, the tools that will be required for effective

conservation in the future extend well beyond traditional

studies in identification, species biology, and ecology.

Rather, they include areas such as conservation genetics,

eDNA, landscape ecology, hydrology, and restoration

engineering. Several conservation projects have already

benefitted from the combined expertise of scientists from

diverse backgrounds working together on mollusk conser-

vation. Examples include key contributions of hydrologists

and engineers in dam removal, water management, and

riparian restoration projects; the critical work of chemists

and toxicologists in deriving water quality standards that are

protective of mollusks; and the foundational products of

social scientists and climatologists in determining how

climate-driven land-use changes may affect the distribution

of mollusks on the landscape. We should continue to

promote this multi-disciplinary approach to mollusk conser-

vation.

Issue 9 – Seek consistent, long-term funding to supportmollusk conservation efforts.

Goal: Increase funding for mollusk conservation.

Strategies:

1. Identify and create a database of existing sources of

funding for mollusk research and management activities.

2. Take advantage of existing local, state, national, and

international grant programs wherever possible.

3. Advocate for prioritization of existing funding to mollusk

research and management.

4. Develop guidelines for monetary mitigation banking.

Overview

Most of the funding for mollusk conservation has come

from federal resource agencies, state wildlife agencies,

universities, regulatory agencies, private conservation organi-

zations, mitigation funds, and damage assessments under the

Clean Water Act and the Comprehensive Environmental

Response, Compensation, and Liability Act (CERCLA).

Funding comes in many forms, including funding to support

staff in organizations that are charged with restoration or

protection of mollusk species and communities. However,

federal and state budgets have been cut in recent years, and

private funding for mollusk conservation and restoration by

way of settlements is ironically driven by catastrophic events

(e.g., chemical spills) rather than by strategic allocations of

funds.

Success stories in funding mollusk conservation

Some mitigation funds have targeted activities that benefit

mussels, such as the Mussel Mitigation Trust Fund (estab-

lished to mitigate for impacts from an Ohio River power plant)

and the National Fish and Wildlife Foundation’s Freshwater

Mussel Conservation Fund (settlement with Tennessee Shell

Company). Most of these funds are short-lived; when their

intended purposes end and expenditures are depleted, there is

little funding for long-term monitoring. Also, there have been

millions of dollars in settlements or court awards from natural

resource damage claims under CERCLA. Both the mitigation

funds and settlement awards have supported key research and

management activities in mollusk conservation far beyond the

geographic scope of the aquatic systems damaged.

There are some examples of consistent and recurring

sources of funding. Since 2000, the SWG program has

provided federal funds to develop and implement programs

that benefit wildlife and their habitats, especially non-game

species. Priority is placed on projects that benefit species of

greatest conservation need, and many states have identified

mollusks as a focal group. These funds must be used to

address conservation needs identified within a state’s CWAP

such as research, surveys, species and habitat management, or

monitoring. In turn, the states administer funds to undertake

work on their own or support cooperator projects. Since 2008,

15NATIONAL STRATEGY FOR FRESHWATER MOLLUSKS

Congress has authorized funding for a competitive SWG

program to encourage multi-partner projects that implement

actions contained in the state CWAP. There is substantial

funding for these programs; in 2014, there was $45.7 million

for all states and territories and another $8.6 million through

the competitive grants program. Mollusk conservation also

gets a creative funding ‘boost’ from enlightened fish

ecologists, whose fishery studies yield valuable information

on mollusks.

Since so many mollusks are on the federal list of

endangered or threatened species, the USFWS provides grants

to states and territories for species and habitat conservation on

private lands. This program is known as Section 6 of the ESA

and four categories of grants are funded: Conservation Grants,

Habitat Conservation Plan Land Acquisition, Habitat Conser-

vation Planning Assistance, and Recovery Land Acquisition

Grants. Across the country, this grant program provides a

substantial amount of money (e.g., $35 million in 2014) to

support species conservation efforts. Although cost-sharing is

required, the percentages are small, ranging from 10–25%.

Opportunities for funding mollusk conservation

The need for long term, consistent funding overlays all

other issues in mollusk conservation. Without stable, adequate

funding, long-term research and management actions cannot

be implemented and evaluated. Mitigation funding on larger

geographic scales is emerging (e.g., NiSource Habitat

Conservation Plan) and some of these projects have the

potential to benefit mollusk conservation. We should evaluate

the costs and benefits of managing a landscape-level

mitigation bank and a mollusk conservation trust fund. States

can increase their chances of securing competitive SWGs by

collaborating and developing multi-state, multi-species, and

cross-regional proposals as well as working across disciplines.

States that do not currently consider mollusks should add them

to their imperilment lists, where warranted.

There is also the option of advocating for re-programming

of existing conservation funding to support mollusk activities;

but with shrinking allocations in federal and state budgets, and

strong competition for available funding, it may be a losing

proposition. For example, there are not enough resources (e.g.,

funding, staff) currently allocated to recover all endangered

mollusks, let alone keep common species common. There are

numerous watershed groups operating on local and regional

scales that can secure grants to work on stream restoration and

concurrent mollusk recovery (e.g., Friends of the Clinch,

Black Diamond, Upper Tennessee River Roundtable). If the

conservation community can continue to convey the message

that healthy mollusk populations are good for rivers and lakes,

and therefore good for fishing, there is a large constituency of

anglers and other outdoor enthusiasts who may advocate for

funding. Additionally, states and non-profit conservation

organizations are eligible for National Fish Passage and

National Fish Habitat Partnership program grants. By

strategically targeting habitat restoration (e.g., dam removal,

culvert replacement) that benefits both fish and mollusks, these

programs offer great opportunities to fund projects that benefit

aquatic fauna over the long-term. Potential new sources of

funding can be hard to predict, given the fast changing

complexion of private and public funding mechanisms. The

FMCS can play a key role in advocating for new and unique

opportunities and centralizing information on potential sources

of funding by creating and maintaining a database of those

sources.

Issue 10 – Coordinate a national strategy for theconservation of mollusk resources.

Goal: Increase coordination and information sharing

among local, state, regional, national, and international

partners in conserving mollusk resources.

Strategies:

1. Establish an ad hoc committee every 12 years to review and

update the National Strategy.

2. Revise the National Strategy document every 15 years and

implement it at multiple scales.

3. Help integrate the national strategies into regional,

ecosystem, and state action plans.

4. Increase collaboration with international partners.

5. Encourage publication of research and management

actions.

As the human population continues to increase and threats

affecting mollusks and their habitats continue to evolve,

researchers and managers will be challenged to meet all of the

conservation needs of mollusks. Therefore, effective conser-

vation of mollusks in the future will require coordinated efforts

from a larger community of partners and stakeholders. Our

recommendations for the future include suggested areas of

urgently needed research, improved taxonomic inclusiveness,

global representation, and regular updates of the National

Strategy. The current topics that need research to conserve and

restore mollusks have been described in Issues 1 to 9.

Coordination of research information and activities will be

imperative for the conservation of mollusks. The information

in this National Strategy should be integrated into ecosystem

and state, regional, and international action plans.

This National Strategy, while greater in breadth taxonom-

ically than the 1998 Strategy, excludes a group of bivalve

mollusks of concern, the Sphaeriidae. This group remains too

poorly known for inclusion at this time. We hope this

document spurs research on this group of mollusks to allow

their inclusion in the next National Strategy.

The 1998 National Strategy greatly improved coordination

across the mollusk community, agencies and organizations,

academia, and private citizens. We anticipate that this

document will be similarly used at local, regional, national,

and international levels to help researchers and managers

prioritize the needed activities to conserve and restore mollusk

16 FMCS 2016

assemblages. A regularly revised National Strategy will serve

as guide for decision-makers that prioritize research funding

and activities, especially as new information becomes

available and new stressors emerge.

Because declines in mollusk faunas are worldwide,

research collaboration across aquatic systems, both nationally

and internationally, will provide insights into mollusk biology

and ecology that might not be obvious when addressed at only

a local or regional level. Outreach and coordination activities

among continental-based groups and the global community is,

however, in its infancy and we encourage the mollusk

community to look globally in their conservation efforts.

Future revisions to the National Strategy should broaden its

geographic scope, encourage international collaboration, and

broaden the taxonomic range of mollusks considered.

ACKNOWLEDGMENTS

The authors thank the membership of the FMCS who

suggested and approved the issues described in this manu-

script. We thank Emy Monroe, Rita Villella, and Dave Zanatta

for contributions to this manuscript. We thank Jeremy

Tiemann, Russell Minton, John Jenkinson, Rachael Hoch,

Lyubov Burlakova, and 3 anonymous reviewers for critical

reviews of this manuscript. Any use of trade, product, or firm

names is for descriptive purposes only and does not imply

endorsement by the U.S. Government. The views expressed in

this article are the authors and do not necessarily represent

those of the U.S. Department of Interior.

REFERENCES

Allen, D. C., and C. C. Vaughn. 2011. Density-dependent biodiversity effects

on physical habitat modification by freshwater bivalves. Ecology

92:1013–1019.

Allen, D. C., C. C. Vaughn, J. F. Kelly, J. T. Cooper, and M. H. Engel. 2012.

Bottom-up biodiversity effects increase resource subsidy flux between

ecosystems. Ecology 93:2165–2174.

Angelini, C., A. H. Altieri, B. R. Silliman, and M. D. Bertness. 2011.

Interactions among foundation species and their consequences for

community organization, biodiversity, and conservation. Bioscience

61:782–789.

Archambault, J. M., W. G. Cope, and T. J. Kwak. 2013. Burrowing, byssus,

and biomarkers: behavioral and physiological indicators of sublethal

thermal stress in freshwater mussels (Unionidae). Marine and Freshwater

Behaviour and Physiology 46:229–250.

Archambault, J. M., W. G. Cope, and T. J. Kwak. 2014. Survival and behavior

of juvenile unionid mussels exposed to thermal stress and dewatering in

the presence of a thermal gradient. Freshwater Biology 59:601–613.

Arthington, A. H., S. E. Bunn, N. L. Poff, and R. J. Naiman. 2006. The

challenge of providing environmental flow rules to sustain river

ecosystems. Ecological Applications 16:1311–1318.

Atkinson, C. L., A. D. Christian, D. E. Spooner, and C. C. Vaughn. 2014a.

Long-lived organisms provide an integrative footprint of agricultural land

use. Ecological Applications 24:375–384.

Atkinson, C. L., J. P. Julian, and C. C. Vaughn. 2014b. Species and function

lost: role of drought in structuring stream communities. Biological

Conservation 176:30–38.

Atkinson, C. L., J. F. Kelly, and C. C. Vaughn. 2014c. Tracing consumer-

derived nitrogen in riverine food webs. Ecosystems 17:485–496.

Atkinson, C. L., and C. C. Vaughn. 2015. Biogeochemical hotspots: temporal

and spatial scaling of the impact of freshwater mussels on ecosystem

function. Freshwater Biology 60:563–574.

Atkinson, C. L., C. C. Vaughn, K. J. Forshay, and J. T. Cooper. 2013.

Aggregated filter-feeding consumers alter nutrient limitation: consequenc-

es for ecosystem and community dynamics. Ecology 94:1359–1369.

Augspurger, T., F. J. Dwyer, C. G. Ingersoll, and C. M. Kane. 2007. Advances

and opportunities in assessing the contaminant sensitivity of freshwater

mussel early life stages. Environmental Toxicology and Chemistry

26:2025–2028.

Augspurger, T., A. E. Keller, M. C. Black, W. G. Cope, and F. J. Dwyer. 2003.

Water quality guidance for protection of freshwater mussels (Unionidae)

from ammonia exposure. Environmental Toxicology and Chemistry

22:2569–2575.

Barnhart, M. C. 2002. Propagation and culture of mussel species of special

concern. Unpublished report, U.S. Fish and Wildlife Service and Missouri

Department of Conservation, Columbia, Missouri. Available at: http://

www.researchgate.net/publication/271074461_Propagation_and_

Culture_of_Mussel_Species_of_Concern_Annual_Report_for_2002 (ac-

cessed December 1, 2015).

Barnhart, M. C. 2006. Buckets of muckets: a compact system for rearing

juvenile freshwater mussels. Aquaculture 254:227–233.

Barnhart, M. C. 2010. Unio gallery. Available at: http://unionid.missouristate.

edu (accessed December 1, 2015).

Barnhart, M. C., W. R. Haag, and W. N. Roston. 2008. Adaptations to host

infection and larval parasitism in Unionoida. Journal of the North

American Benthological Society 27:370–394.

Beck, M. W., R. D. Brumbaugh, L. Airoldi, A. Carranza, L. D. Coen, C.

Crawford, O. Defeo, G. J. Edgar, B. Hancock, M. C. Kay, H. S. Lenihan,

M. W. Luckenbach, C. L. Toropova, G. Zhang, and X. Guo. 2011. Oyster

reefs at risk and recommendations for conservation, restoration, and

management. BioScience 61:107–116.

Bogan, A. E., and K. J. Roe. 2008. Freshwater bivalve (Unioniformes)

diversity, systematics, and evolution: status and future directions. Journal

of the North American Benthological Society 27:349–369.

Brauman, K. A., G. C. Daily, T. K. Duarte, and H. A. Mooney. 2007. The

nature and value of ecosystem services: an overview highlighting

hydrologic services. The Annual Review of Environment and Resources

32:6.1–6.32.

Brim Box, J., R. M. Dorazio, and W. D. Liddell. 2002. Relationships between

streambed substrate characteristics and freshwater mussels (Bivalvia:

Unionidae) in coastal plain streams. Journal of the North American

Benthological Society 21:253–260.

Brim Box, J., J. Howard, D. Wolf, C. O’Brien, D. Nez, and D. Close. 2006.

Freshwater mussels (Bivalvia: Unionoida) of the Umatilla and Middle

Fork John Day Rivers in Eastern Oregon. Northwest Science 80:95–107.

Brown, K. M., B. Lang, and K. E. Perez. 2008. The conservation ecology of

North American pleurocerid and hydrobiid gastropods. Journal of the

North American Benthological Society 27:484–495.

Bruesewitz, D. A., J. L. Tank, and S. K. Hamilton. 2009. Seasonal effects of

zebra mussels on littoral nitrogen transformation rates in Gull Lake,

Michigan. Freshwater Biology 54:1427–1443.

Buck, E. H., M. L. Corn, K. Alexander, P. A. Sheikh, and R. Meltz. 2012. The

Endangered Species Act (ESA) in the 112th Congress: conflicting values

and difficult choices. Congressional Research Service. Report No.

R411608. Available at: http://www.fas.org/sgp/crs/misc/R41608.pdf (ac-

cessed December 1, 2015).

Burlakova, L. E., and V. A. Karatayev. 2010. State-wide assessment of

unionid diversity in Texas. Unpublished report, Texas Parks and Wildlife,

Austin. Available at: https://tpwd.texas.gov/huntwild/wild/wildlife_

diversity/nongame/mussels/media/burlakova-statewide-assessment-of-

unionid-diversity-in-texas1.pdf (accessed December 1, 2015).

17NATIONAL STRATEGY FOR FRESHWATER MOLLUSKS

Burlakova, L. E., A. Y. Karatayev, V. A. Karatayev, M. E. May, D. L.

Bennett, and M. J. Cook. 2011. Endemic species: contribution to

community uniqueness, effect of habitat alteration, and conservation

priorities. Biological Conservation 144:155–165.

Butler, R. S. 2007. Status assessment report for the Snuffbox, Epioblasma

triquetra, a freshwater mussel occurring in the Mississippi River and

Great Lakes Basins. Unpublished report, U.S. Fish and Wildlife Service,

Asheville, North Carolina. Available at: https://www.researchgate.net/

profile/Robert_Butler5/publication/275651716_Status_Assessment_

Report_for_the_Snuffbox_Epioblasma_triquetra_a_Freshwater_Mussel_

Occurring_in_the_Mississippi_River_and_Great_Lakes_Basins/links/

554295aa0cf234bdb21a173e.pdf (accessed December 1, 2015).

Calabro, E. J., B. A. Murry, D. A. Woolnough, and D. G. Uzarski. 2013.

Application and transferability of Great Lakes coastal wetland indices of

biotic integrity to high quality inland lakes of Beaver Island in northern

Lake Michigan. Aquatic Ecosystem Health and Management 16:338–346.

Cardinale, B. J., M. A. Palmer, and S. L. Collins. 2002. Species diversity

enhances ecosystem functioning through interspecific facilitation. Nature

415:426–429.

Carey, C. S., J. W. Jones, R. S. Butler, and E. M. Hallerman. 2015. Restoring

the endangered oyster mussel (Epioblasma capsaeformis) to the upper

Clinch River, Virginia: an evaluation of population restoration techniques.

Restoration Ecology 23:447–454.

Chong, J. P., J. C. Brim Box, J. K. Howard, D. Wolf, T. L. Meyers, and K. E.

Mock. 2008. Three deeply divided lineages of the freshwater mussel

genus Anodonta in Western North America. Conservation Genetics

9:1303–1309.

Coogan, M. A., and T. W. La Point. 2008. Snail bioaccumulation of

triclocarban, triclosan, and methyltriclosan in a North Texas, USA, stream

affected by wastewater treatment plant runoff. Environmental Toxicology

and Chemistry 27:1788–1793.

Cope, W. G., T. J. Newton, and C. M. Gatenby. 2003. Review of techniques to

prevent introduction of zebra mussels (Driessena polymorpha) during

native mussel (Unionidea) conservation activities. Journal of Shellfish

Research 22:177–184.

Crabtree, D. L., and T. A. Smith. 2009. Population attributes of an endangered

mussel, Epioblasma torulosa rangiana (Northern Riffleshell), in French

Creek and implications for its recovery. Northeastern Naturalist 16:339–

354.

Crail, T. D., R. A. Krebs, and D. T. Zanatta. 2011. Unionid mussels from

nearshore zones of Lake Erie. Journal of Great Lakes Research 37:199–

202.

CRMRC (Cumberlandian Region Mollusk Restoration Committee). 2010.

Plan for the population restoration and conservation of freshwater

mollusks of the Cumberlandian Region. Unpublished report, Asheville,

North Carolina. Available from Robert S. Butler, U.S. Fish and Wildlife

Service, 160 Zillicoa Street, Asheville, North Carolina 28801 USA.

Davis, M. 2005. Clam chronicles: an account of activities associated with

efforts to propagate and repatriate Lampsilis higginsii in the Mississippi

River, Minnesota. Unpublished report, Minnesota Department of Natural

Resources, Lake City. Available at: http://www.fws.gov/midwest/mussel/

documents/clam_chronicles_2005.pdf (accessed December 1, 2015).

Dillon, R. T., Jr., M. Ashton, M. Kohl, W. Reeves, T. Smith, T. Stewart, and

B. Watson. 2013. The freshwater gastropods of North America. Available

at: http://www.fwgna.org (accessed December 1, 2015).

Dodds, W. K., J. S. Perkin, and J. E. Gerken. 2013. Human impact on

freshwater ecosystem services: a global perspective. Environmental

Science and Technology 47:9061–9068.

Downing, J. A., P. Van Meter, and D. A. Woolnough. 2010. Suspects and

evidence: a review of the causes of decline and extirpation in freshwater

mussels. Animal Biodiversity and Conservation 33:151–185.

Dunca, E., B. R. Schone, and H. Mutvei. 2005. Freshwater bivalves tell of past

climates: but how clearly do shells from polluted rivers speak?

Palaeogeography, Paleaoclimatology, Palaeoecology 228:43–57.

Dunn, H., S. Zigler, J. Duyvejonck, and T. Newton. 2012. Methods to assess

and monitor mussel communities in the Upper Mississippi River system.

Unpublished report, U.S. Army Corps of Engineers, Rock Island, Illinois.

Available from Teresa J. Newton, U.S. Geological Survey, 2630 Fanta

Reed Road, La Crosse, WI 54603 USA.

Evans, R. R., and S. J. Ray. 2010. Distribution and environmental influences

on freshwater gastropods from lotic systems and springs in Pennsylvania,

USA, with conservation recommendations. American Malacological

Bulletin 28:135–150.

Galbraith, H. S., D. E. Spooner, and C. C. Vaughn. 2010. Synergistic effects of

regional climate patterns and local water management on freshwater

mussel communities. Biological Conservation 143:1175–1183.

Gangloff, M. M., and J. W. Feminella. 2007. Stream geomorphology

influences mussel abundance in Southern Appalachian streams, U.S.A.

Freshwater Biology 52:64–74.

Gangloff, M. M., E. E. Hartfield, D. C. Werneke, and J. W. Feminella. 2011.

Associations between small dams and mollusk assemblages in Alabama

streams. Journal of the North American Benthological Society 30:1107–

1116.

Ganser, A. G., T. J. Newton, and R. J. Haro. 2013. The effects of elevated

water temperatures on native juvenile mussels: implications for the effects

of climate change. Freshwater Science 32:1168–1177.

Gatenby, C. M., D. A. Kreeger, M. A. Patterson, M. Marinni, and R. J. Neves.

2013. Clearance rates of Villosa iris (Bivalvia: Unionidae) fed different

rations of the alga Neochloris oleoabundans. Walkerana 16:9–20.

Gatenby, C. M., P. A. Morrison, R. J. Neves, and B. C. Parker. 2000. A

protocol for the salvage and quarantine of unionid mussels from zebra

mussel-infested waters. Pages 9–18 in R. A. Tankersley, D. I. Warmolts,

G. T. Watters, B. J. Armitage, P. D. Johnson, and R. S. Butler, editors.

Part I. Proceedings of the conservation, captive care, and propagation of

freshwater mussels symposium. Ohio Biological Survey Special Publica-

tion, Columbus.

Gates, K. K., C. C. Vaughn, and J. P. Julian. 2015. Developing environmental

flow recommendations for freshwater mussels using the biological traits of

species guilds. Freshwater Biology 60:620–635.

Geist, J. 2010. Strategies for the conservation of endangered freshwater pearl

mussels (Margaritifera margaritifera L.): a synthesis of conservation

genetics and ecology. Hydrobiologia 644:69–88.

Gillis, P. 2011. Assessing the toxicity of sodium chloride to the glochidia of

freshwater mussels: implications for salinization of surface waters.

Environmental Pollution 159:1702–1708.

Goldberg, C. S., A. Sepulveda, A. Ray, J. Baumgardt, and L. P. Waits. 2013.

Environmental DNA as a new method for early detection of New Zealand

mudsnails (Potamopyrgus antipodarum). Freshwater Science 32:792–

800.

Gough, H. M., A. M. Gascho Landis, and J. A. Stoeckel. 2012. Behaviour and

physiology are linked in the responses of freshwater mussels to drought.

Freshwater Biology 57:2356–2366.

Graf, D. L. 2001. The cleansing of the Augean stables: or a lexicon of the

nominal species of the Pleuroceridae (Gastropoda: Prosobranchia) of

recent North America, north of Mexico. Walkerana 12:1–124.

Graf, D. L. 2013. Patterns of freshwater bivalve global diversity and the state

of phylogenetic studies on the Unionoida, Sphaeriidae, and Cyrenidae.

American Malacological Bulletin 31:135–153.

Graf, D. L., and K. S. Cummings. 2007. Review of the systematics and global

diversity of freshwater mussel species (Bivalvia: Unionoida). Journal of

Molluscan Studies 73:291–314.

Graf, D. L., and K. S. Cummings. 2015. The MUSSEL Project. Available at:

http://mussel-project.uwsp.edu/ (accessed December 1, 2015).

Gray, M. W., and D. Kreeger. 2014. Monitoring fitness of caged mussels

(Elliptio complanata) to assess and prioritize streams for restoration.

Aquatic Conservation: Marine and Freshwater Ecosystems 24:218–230.

Greene, H. 2005. Organisms in nature as a central focus in biology. Trends in

Ecology and Evolution 20:23–27.

18 FMCS 2016

Haag, W. R. 2009. Past and future patterns of freshwater mussel extinctions in

North America during the Holocene. Pages 107–128 in S. Turvey, editor.

Holocene Extinctions. Oxford University Press, New York.

Haag, W. R. 2012. North American freshwater mussels: natural history,

ecology, and conservation. Cambridge University Press, Cambridge,

United Kingdom.

Haag, W. R., and M. L. Warren, Jr. 2008. Effects of severe drought on

freshwater mussel assemblages. Transactions of the American Fisheries

Society 137:1165–1178.

Haag, W. R., and M. L. Warren, Jr. 2010. Diversity, abundance, and size

structure of bivalve assemblages in the Sipsey River, Alabama. Aquatic

Conservation: Marine and Freshwater Ecosystems 20:655–667.

Haag, W. R., and J. D. Williams. 2014. Biodiversity on the brink: an

assessment of conservation strategies for North American freshwater

mussels. Hydrobiologia 735:45–60.

Hall, R. O., J. L. Tank, and M. F. Dybdahl. 2003. Exotic snails dominate

nitrogen and carbon cycling in a highly productive stream. Frontiers in

Ecology and the Environment 1:407–411.

Hall, R. O., M. F. Dybdahl, and M. C. VanderLoop. 2006. Extremely high

secondary production of introduced snails in rivers. Ecological Applica-

tions 16:1121–1131.

Harried, B., K. Fischer, K. E. Perez, and G. J. Sandland. 2015. Assessing

infection patterns in Chinese mystery snails from Wisconsin, USA, using

field and laboratory approaches. Aquatic Invasions 10:169–175.

Hartfield, P. D. 2003. Freshwater mussels and snails of the Mobile River basin:

plan for controlled propagation, augmentation, and reintroduction.

Unpublished report, U.S. Fish and Wildlife Service, Jackson, Mississippi.

Available from: Paul D. Hartfield, U.S. Fish and Wildlife Service, 6578

Dogwood View Parkway, Suite A, Jackson, Mississippi 39213 USA.

Hershler, R., H.-P. Liu, and D. L. Gustafson. 2008. A second species of

Pyrgulopsis (Hydrobiidae) from the Missouri River Basin, with molecular

evidence supporting faunal origin through Pliocene stream capture across

the Northern Continental Divide. Journal of Molluscan Studies 74:403–

413.

Hershler, R., and H.-P. Liu. 2009. New species and records of Pyrgulopsis

(Gastropoda: Hydrobiidae) from the Snake River basin, Southeastern

Oregon: further delineation of a highly imperiled fauna. Zootaxa 2006:1–

22.

Hershler, R., and H.-P. Liu. 2010. Two new, possibly threatened species of

Pyrgulopsis (Gastropoda: Hydrobiidae) from Southwestern California.

Zootaxa 2343:1–17.

Hershler, R., J. M. Pierson, and R. S. Krotzer. 1990. Rediscovery of Tulotoma

magnifica (Conrad) (Gastropoda: Viviparidae). Proceedings of the

Biological Society of Washington 103:815–824.

Hill, W. R. 1992. Food limitation and interspecific competition in snail-

dominated streams. Canadian Journal of Fisheries and Aquatic Sciences

49:1257–1267.

Hoftyzer, E., J. D. Ackerman, T. J. Morris, and G. L. Mackie. 2008. Genetic

and environmental implications of reintroducing laboratory-raised unionid

mussels to the wild. Canadian Journal of Fisheries and Aquatic Sciences

65:1217–1229.

Holznagel, W. E., and C. Lydeard. 2000. A molecular phylogeny of North

American Pleuroceridae (Gastropoda: Cerithioidea) based on mitochon-

drial 16S rDNA sequences. Journal of Molluscan Studies 66:233–257.

Huryn, A. D., J. W. Koebel, and A. C. Benke. 1994. Life history and longevity

of the pleurocerid snail Elimia: a comparative study of eight populations.

Journal of the North American Benthological Society 13:540–556.

Johnson, P. D., A. E. Bogan, K. M. Brown, N. M. Burkhead, J. R. Cordeiro, J.

T. Garner, P. D. Hartfield, D. A. W. Lepitzki, G. L. Mackie, E. Pip, T. A.

Tarpley, J. S. Tiemann, N. V. Whelan, and E. E. Strong. 2013.

Conservation status of freshwater gastropods of Canada and the United

States. Fisheries 38:247–282.

Johnson, P. T. J., J. D. Olden, C. T. Solomon, and M. J. Vander Zanden. 2009.

Interactions among invaders: community and ecosystem effects of

multiple invasive species in an experimental aquatic system. Oecologia

159:161–170.

Jones, J. W., E. M. Hallerman, and R. J. Neves. 2006. Genetic management

guidelines for captive propagation of freshwater mussels (Unionoidea).

Journal of Shellfish Research 25:527–535.

Jones, J. W., and R. J. Neves. 2010. Descriptions of a new species and a new

subspecies of freshwater mussels, Epioblasma ahlstedti and Epioblasma

florentina aureola (Bivalvia: Unionidae), in the Tennessee River drainage,

USA. The Nautilus 124:77–92.

Jones, J. W., R. J. Neves, S. A. Ahlstedt, and E. M. Hallerman. 2006. A

holistic approach to taxonomic evaluation of two closely related

endangered freshwater mussel species, the oyster mussel Epioblasma

capsaeformis and tan riffleshell Epioblasma florentina walkeri (Bivalvia:

Unionidae). Journal of Molluscan Studies 72:267–283.

Jones, J. W., R. J. Neves, and E. M. Hallerman. 2012. Population performance

criteria to evaluate reintroduction and recovery of two endangered mussel

species, Epioblasma brevidens and Epioblasma capsaeformis (Bivalvia:

Unionidae). Walkerana 15:27–44.

Jones, J. W., R. J. Neves, and E. M. Hallerman. 2015. Historical demography

of freshwater mussels (Bivalvia: Unionidae): genetic evidence for

population expansion and contraction during the late Pleistocene and

Holocene. Biological Journal of the Linnean Society 114:376–397.

Karatayev, A. Y., L. E. Burlakova, S. E. Mastitsky, and D. K. Padilla. 2015.

Predicting the spread of aquatic invaders: insight from 200 years of

invasion by zebra mussels. Ecological Applications 25:430–440.

Kellogg, M. L., J. C. Cornwell, M. S. Owens, and K. T. Paynter. 2013.

Denitrification and nutrient assimilation on a restored oyster reef. Marine

Ecology Progress Series 480:1–19.

Klippel, W. E., and D. F. Morey. 1986. Contextual and nutritional analysis of

freshwater gastropods from middle Archaic deposits at the Hayes Site,

middle Tennessee. American Antiquity 51:799–813.

Kurth, J., C. Loftin, J. Zydlewski, and J. Rhymer. 2007. PIT tags increase

effectiveness of freshwater mussel recaptures. Journal of the North

American Benthological Society 26:253–260.

Lande, R., S. Engen, and B.-E. Sæther. 2003. Stochastic population dynamics

in ecology and conservation. Oxford University Press, Oxford, United

Kingdom.

Lane, T., H. Dan, and J. Jones. 2014. Reintroduction of endangered freshwater

mussel populations to high priority geographic areas in the upper

Tennessee River system. Unpublished report, U.S. Fish and Wildlife

Service, Asheville, North Carolina. Available from: Robert S. Butler, U.S.

Fish and Wildlife Service, 160 Zillicoa Street, Asheville, North Carolina

28801 USA.

Lange, K., C. R. Townsend, and C. D. Matthaei. 2014. Can biological traits of

stream invertebrates help disentangle the effects of multiple stressors in an

agriculural catchment? Freshwater Biology 59:2431–2446.

Lydeard, C., R. H. Cowie, W. F. Ponder, A. E. Bogan, P. Bouchet, S. A. Clark,

K. S. Cummings, T. J. Frest, P. Gargominy, K. E. Perez, B. Roth, M.

Seddon, E. E. Strong, and F. G. Thompson. 2004. The global decline of

nonmarine mollusks. BioScience 54:321–330.

Lydeard, C., W. E. Holznagel, J. Garner, P. Hartfield, and J. M. Pierson. 1997.

A molecular phylogeny of Mobile River drainage basin pleurocerid snails

(Caenogastropoda: Cerithioidea). Molecular Phylogenetics and Evolution

7:117–128.

Lysne, S. J., K. E. Perez, K. W. Brown, R. L. Minton, and J. D. Sides. 2008. A

review of freshwater gastropod conservation: challenges and opportuni-

ties. Journal of the North American Benthological Society 27:463–470.

McMurray, S.E. 2015. Programmatic guidance for the controlled propagation,

augmentation, and reintroduction of freshwater mussels in Missouri.

Missouri Department of Conservation, Columbia, Missouri. Available

from Steve McMurray, Missouri Department of Conservation, 3500 East

Gans Road, Columbia, MO 65201-8992 USA.

Mair, R. A. 2013. A suitable diet and culture system for rearing freshwater

mussels at White Sulphur Springs National Fish Hatchery, West Virginia.

19NATIONAL STRATEGY FOR FRESHWATER MOLLUSKS

Master’s thesis, Virginia Polytechnic Institute and State University,

Blacksburg.

May, R. M. 1990. Taxonomy as destiny. Nature 347:129–130.

Menzel, S., and J. Teng. 2010. Ecosystem services as a stakeholder-driven

concept for conservation science. Conservation Biology 24:907–909.

Minton, R. L., and C. Lydeard. 2003. Phylogeny, taxonomy, genetics and

global heritage ranks of an imperiled, freshwater snail genus Lithasia

(Pleuroceridae). Molecular Ecology 12:75–87.

Morningstar, C., K. Inoue, M. Sei, B. K. Lang, and D. J. Berg. 2014.

Quantifying morphological and genetic variation of sympatric populations

to guide conservation of endangered, micro-endemic springsnails. Aquatic

Conservation: Marine and Freshwater Ecosystems 24:536–545.

Morrison, P. A. 2012. 2012 accomplishment report, Ohio River restoration

project, NRDA Project 0237. Unpublished report, U.S. Fish and Wildlife

Service, Williamstown, West Virginia. Available from: Patricia A.

Morrison, U.S. Fish and Wildlife Service, Ohio River Islands National

Wildlife Refuge, 3982 Waverly Road, Williamstown, West Virginia

26187 USA.

Morrison, P., C. Gatenby, J. Devers, M. Patterson, and R. Mair. 2013. Salvage

and refuge of target mussel species (2004–2012): mitigation of bridge

replacement at East Brady, Pennsylvania. Unpublished report, U.S. Fish

and Wildlife Service, Williamstown, West Virginia. Available from:

Patricia A. Morrison, U.S. Fish and Wildlife Service, Ohio River Islands

National Wildlife Refuge, 3982 Waverly Road, Williamstown, West

Virginia 26187 USA.

Naimo, T. J. 1995. A review of the effects of heavy metals on freshwater

mussels. Ecotoxicology 4:341–362.

Nalepa, T. F., and D. W. Schloesser. 2014. Quagga and zebra mussels:

biology, impacts, and control. CRC Press, Taylor & Francis, Boca Raton,

Florida.

NatureServe. 2015. NatureServe, a network connecting science with

conservation. Available at: http://www.natureserve.org (accessed Decem-

ber 1, 2015).

Neves, R. J. 2004. Propagation of endangered freshwater mussels in North

America. Journal of Conchology Special Publication 3:69–80.

Newton, T. J., J. W. Allran, J. A. O’Donnell, M. R. Bartsch, and W. B.

Richardson. 2003. Effects of ammonia on juvenile unionid mussels

(Lampsilis cardium) in laboratory sediment toxicity tests. Environmental

Toxicology and Chemistry 22:2554–2560.

Newton, T. J., and M. R. Bartsch. 2007. Lethal and sublethal effects of

ammonia to juvenile Lampsilis mussels (Unionidae) in sediment and

water-only exposures. Environmental Toxicology and Chemistry

26:2057–2065.

Newton, T. J., and W. G. Cope. 2007. Biomarker responses of unionid mussels

to environmental contaminants. Pages 257–284 in J. L. Farris and J. H.

Van Hassel, editors. Freshwater Bivalve Ecotoxicology. SETAC Press,

Pensacola, Florida, and Taylor & Francis, Boca Raton, Florida.

Newton, T. J., C. C. Vaughn, D. E. Spooner, S. J. Nichols, and M. T. Arts.

2013. Profiles of biochemical tracers in unionid mussels across a broad

geographic range. Journal of Shellfish Research 32:497–507.

Newton, T. J., D. A. Woolnough, and D. L. Strayer. 2008. Using landscape

ecology to understand freshwater mussel populations. Journal of the North

American Benthological Society 27:424–439.

Newton, T. J., S. J. Zigler, J. T. Rogala, B. R. Gray, and M. Davis. 2011.

Population assessment and potential functional roles of native mussels in

the Upper Mississippi River. Aquatic Conservation: Marine and

Freshwater Ecosystems 21:122–131.

Newton, T. J., S. J. Zigler, and B. R. Gray. 2015. Mortality, movement, and

behavior of native mussels during a planned water-level drawdown in the

Upper Mississippi River. Freshwater Biology 60:1–15.

Nichols, S. J., and D. A. Wilcox. 1997. Burrowing saves Lake Erie clams.

Nature 389:921.

Nijhuis, M. 2009. The Cahaba: a river of riches. Available at: http://www.

smithsonianmag.com/science-nature/the-cahaba-a-river-of-riches-

34214889/?no-ist (accessed December 1, 2015).

NNMCC (National Native Mussel Conservation Committee). 1998. National

strategy for the conservation of native freshwater mussels. Journal of

Shellfish Research 17:1419–1428.

Olden, J. D., M. J. Kennard, J. J. Lawler, and N. L. Poff. 2010. Challenges and

opportunities in implementing managed relocation for conservation of

freshwater species. Conservation Biology 25:40–47.

Olson, Z. H., J. T. Briggler, and R. N. Williams. 2012. An eDNA approach to

detect eastern hellbenders (Crytobranchus a. alleganiensis) using samples

of water. Wildlife Research 39:629–636.

Pandolfo, T. J., W. G. Cope, C. Arellano, R. B. Bringolf, M. C. Barnhart, and

D. E. Hammer. 2010. Upper thermal tolerances of early life stages of

freshwater mussels. Journal of the North American Benthological Society

29:959–969.

Pandolfo, T. J., T. J. Kwak, and W. G. Cope. 2012. Thermal tolerances of

freshwater mussels and their host fish: species interactions in a changing

climate. Walkerana: The Journal of the Freshwater Mollusk Conservation

Society 15:69–82.

Perez, K. E., and R. L. Minton. 2008. Practical applications for systematics

and taxonomy in North American freshwater gastropod conservation.

Journal of the North American Benthological Society 27:471–483.

Perez, K. E., and G. Sandland. 2015. Key to Wisconsin freshwater snails.

Available at: http://northamericanlandsnails.com/WIFreshwaterSnailskey/

wifwsnailkey.html (accessed December 1, 2015).

Pilliod, D. S., C. S. Goldberg, R. S. Arkle, and L. P. Waits. 2013. Estimating

occupancy and abundance of stream amphibians using eDNA from

filtered water samples. Canadian Journal of Fisheries and Aquatic

Sciences 70:1123–1130.

Poff, N. L., J. D. Allan, M. B. Bain, J. R. Karr, K. L. Prestgaard, B. D. Richter,

R. E. Sparks, and J. C. Stomberg. 1997. The natural flow regime: a

paradigm for river conservation and restoration. BioScience 47:769–784.

Poff, N. L., J. D. Olden, D. M. Merrit, and D. M. Pepin. 2007.

Homogenization of regional river dynamics and global biodiversity

implications. Proceedings of the National Academy of Sciences

104:5732–5737.

Richardson, J., and J. Loomis. 2009. The total economic value of threatened,

endangered and rare species: an updated meta-analysis. Ecological

Economics 68:1535–1548.

Richardson, T. D., J. F. Scheiring, and K. M. Brown. 1988. Secondary

production of two lotic snails (Pleuroceridae: Elimia). Journal of the North

American Benthological Society 7:234–241.

Rypel, A. L., W. R. Haag, and R. H. Findlay. 2008. Validation of annual

growth rings in freshwater mussel shells using cross dating. Canadian

Journal of Fisheries and Aquatic Sciences 65:2224–2232.

Seppelt, R., C. F. Dormann, F. V. Eppink, S. Lautenbach, and S. Schmidt.

2011. A quantitative review of ecosystem service studies: approaches,

shortcomings and the road ahead. Journal of Applied Ecology 48:630–

636.

Serb, J. M., J. E. Buhay, and C. Lydeard. 2003. Molecular systematics of the

North American freshwater bivalve genus Quadrula (Unionidae:

Ambleminae) based on mitochondrial ND1 sequences. Molecular

Phylogenetics and Evolution 28:1–11.

Sethi, S. A., A. R. Selle, M. W. Doyle, E. H. Stanley, and H. E. Kitchel. 2004.

Response of unionid mussels to dam removal in Koshkonong Creek,

Wisconsin (USA). Hydrobiologia 525:157–165.

Shea, C. P., J. T. Peterson, M. J. Conroy, and J. M. Wisniewski. 2013.

Evaluating the influence of land use, drought and reach isolation on the

occurrence of freshwater mussel species in the lower Flint River Basin,

Georgia (USA). Freshwater Biology 58:382–395.

Smyth, A. R., N. R. Geraldi, and M. F. Piehler. 2013. Oyster-mediated

benthic-pelagic coupling modifies nitrogen pools and processes. Marine

Ecology Progress Series 493:23–30.

Spooner, D. E., and C. C. Vaughn. 2008. A trait-based approach for species’

20 FMCS 2016

roles in aquatic ecosystems: climate change, community structure, and

material cycling. Oecologia 158:307–317.

Spooner, D. E., and C. C. Vaughn. 2009. Species richness and temperature

influence mussel biomass: a partitioning approach applied to natural

communities. Ecology 90:781–790.

Spooner, D. E., M. A. Xenopoulos, C. Schneider, and D. A. Woolnough.

2011. Coextirpation of host-affiliate relationships in rivers: the role of

climate change, water withdrawal, and host-specificity. Global Change

Biology 17:1720–1732.

Statzner, B., and L. A. Beche. 2010. Can biological invertebrate traits resolve

effects of multiple stressors on running water ecosystems? Freshwater

Biology 55:80–119.

Steuer, J. J., T. J. Newton, and S. J. Zigler. 2008. Use of complex hydraulic

variables to predict the distribution and density of unionids in a side

channel of the Upper Mississippi River. Hydrobiologia 610:67–82.

Strayer, D. L. 2008. Freshwater mussel ecology: a multifactor approach to

distribution and abundance. University of California Press, Berkeley.

Strayer, D. L. 2014. Understanding how nutrient cycles and freshwater

mussels (Unionoida) affect one another. Hydrobiologia 735:277–292.

Strayer, D. L., J. A. Downing, W. R. Haag, T. L. King, J. B. Layzer, T. J.

Newton, and S. J. Nichols. 2004. Changing perspectives on pearly

mussels, North America’s most imperiled animals. BioScience 54:429–

439.

Strayer, D. L., and H. Malcolm. 2007. Submersed vegetation as habitat for

invertebrates in the Hudson River estuary. Estuaries and Coasts 30:253–

264.

Strayer, D. L., and J. Ralley. 1993. Microhabitat use by an assemblage of

stream-dwelling unionaceans (Bivalvia), including two rare species of

Alasmidonta. Journal of the North American Benthological Society

12:247–258.

Strayer, D. L., and D. R. Smith. 2003. A guide to sampling freshwater mussel

populations. American Fisheries Society Monograph 8, Bethesda,

Maryland.

Thompson, F. G. 2004. An identification manual for the freshwater snails of

Florida. Available at: http://www.flmnh.ufl.edu/malacology/fl-snail/

snails1.htm (accessed December 1, 2015).

Turgeon, D. D., J. F. Quinn, Jr., A. E. Bogan, E. V. Coan, F. G. Hochberg, W.

G. Lyons, P. M. Mikkelsen, R. J. Neves, C. F. E. Roper, G. Rosenburg, B.

Roth, A. Scheltema, F. G. Thompson, M. Vecchione, and J. D. Williams.

1998. Common and scientific names of aquatic invertebrates from the

United States and Canada: mollusks, 2nd edition. American Fisheries

Society, Special Publication 26, Bethesda, Maryland.

USEPA (U.S. Environmental Protection Agency). 2013. Aquatic life ambient

water quality criteria for ammonia—freshwater. USEPA, 822-R-13-001,

Washington, D.C.

USFWS (U.S. Fish and Wildlife Service). 2004. Recovery plan for Cumber-

land elktoe (Alasmidonta atropurpurea), oyster mussel (Epioblasma

capsaeformis), Cumberlandian combshell (Epioblasma brevidens), purple

bean (Villosa perpurpurea), and rough rabbitsfoot (Quadrula cylindrica

strigillata). Atlanta, Georgia. Available at: http://ecos.fws.gov/docs/

recovery_plan/040524.pdf (accessed December 1, 2015).

USFWS (U.S. Fish and Wildlife Service). 2010. Scaleshell (Leptodea

leptodon) recovery plan. Fort Snelling, Minnesota. Available at: http://

www.fws.gov/midwest/endangered/clams/ScaleshellRecoveryPlan.html

(accessed December 1, 2015).

USFWS (U.S. Fish and Wildlife Service). 2011. Reclassification of the Tulotoma

snail from endangered to threatened. Available at: http://www.fws.gov/

policy/library/2011/2011-13687.html (accessed December 1, 2015).

USFWS (U.S. Fish and Wildlife Service). 2014. Imperiled aquatic species

conservation strategy for the upper Tennessee River basin. Abingdon,

Virginia. Available at: http://applcc.org/projects/trb/imperiled-aquatic-

species-conservation-strategy/copy_of_imperiled-aquatic-species-

conservation-strategy (accessed December 1, 2015).

Vaughn, C. C. 2010. Biodiversity losses and ecosystem function in

freshwaters: emerging conclusions and research directions. BioScience

60:25–35.

Vaughn, C. C., and C. C. Hakenkamp. 2001. The functional role of burrowing

bivalves in freshwater ecosystems. Freshwater Biology 46:1431–1446.

Vaughn, C. C., S. J. Nichols, and D. E. Spooner. 2008. Community and

foodweb ecology of freshwater mussels. Journal of the North American

Benthological Society 27:409–423.

Villella, R. F., D. R. Smith, and D. P. Lemarie. 2004. Estimating survival and

recruitment in a freshwater mussel population using mark-recapture

techniques. American Midland Naturalist 151:114–133.

Wang, N., C. G. Ingersoll, I. E. Greer, D. K. Hardesty, C. D. Ivey, J. L. Kunz,

W. G. Brumbaugh, F. J. Dwyer, A. D. Roberts, T. Augspurger, C. M.

Kane, R. J. Neves, and M. C. Barnhart. 2007a. Chronic toxicity of copper

and ammonia to juvenile freshwater mussels (Unionidae). Environmental

Toxicology and Chemistry 26:2048–2056.

Wang, N., C. G. Ingersoll, D. K. Hardesty, C. D. Ivey, J. L. Kunz, T. W. May,

F. J. Dwyer, A. D. Roberts, T. Augspurger, C. M. Kane, R. J. Neves, and

M. C. Barnhart. 2007b. Acute toxicity of copper, ammonia, and chlorine

to glochidia and juveniles of freshwater mussels (Unionidae). Environ-

mental Toxicology and Chemistry 26:2036–2047.

Watters, G. T., and C. Byrne. 2015. Division of Molluscs, Department of

Evolution, Ecology, and Organismal Biology, The Ohio State University.

Available at: http://www.biosci.ohio-state.edu/~molluscs/OSUM2/ (ac-

cessed December 1, 2015).

Watters, G. T., M. A. Hoggarth, and D. H. Stansbery. 2009. The freshwater

mussels of Ohio. Ohio State University Press, Columbus.

Wethington, A. R., and R. Guralnick. 2004. Are populations of physids from

different hot springs distinctive lineages? American Malacological

Bulletin 18:135–144.

Wethington, A. R., and C. Lydeard. 2007. A molecular phylogeny of Physidae

(Gastropoda: Basommatophora) based on mitochondrial DNA sequences.

Journal of Molluscan Studies 73:241–257.

Whelan, N. V., and E. E. Strong. 2016. Morphology, molecules, and

taxonomy: extreme incongruence in pleurocerids (Gastropoda, Cerithioi-

dea, Pleuroceridae). Zoological Scripta 45:62–87.

Williams, J. D., A. E. Bogan, and J. T. Garner. 2008. Freshwater mussels of

Alabama and the Mobile Basin of Georgia, Mississippi, and Tennessee.

University of Alabama Press, Tuscaloosa.

Williams, J. D., M. L. Warren, Jr., K. S. Cummings, J. L. Harris, and R. J.

Neves. 1993. Conservation status of freshwater mussels of the United

States and Canada. Fisheries 18:6–22.

Zanatta, D. T., J. M. Bossenbroek, L. E. Burlakova, T. D. Crail, F. de Szalay,

T. A. Griffith, D. Kapusinski, A. Y. Karateyev, R. A. Krebs, E. S. Meyer,

W. L. Paterson, T. J. Prescott, M. T. Rowe, D. W. Schloesser, and M. C.

Walsh. 2015. Distribution of native mussel (Unionidae) assemblages in

coastal Lake Erie, Lake St. Clair, and connecting channels, twenty-five

years after the dreissenid invasion. Northeastern Naturalist 22:223–235.

Zigler, S. J., T. J. Newton, J. J. Steuer, M. R. Bartsch, and J. S. Sauer. 2008.

Importance of physical and hydraulic characteristics to unionid mussels: a

retrospective analysis in a reach of large river. Hydrobiologia 598:343–360.

Zimmerman, G. F., and F. A. de Szalay. 2007. Influence of unionid mussels

(Mollusca: Unionidae) on sediment stability: an artificial stream study.

Fundamental and Applied Limnology 168:299–306.

21NATIONAL STRATEGY FOR FRESHWATER MOLLUSKS

Freshwater Mollusk Biology and Conservation 19:22–28, 2016

� Freshwater Mollusk Conservation Society 2016

ARTICLE

INVESTIGATION OF FRESHWATER MUSSEL GLOCHIDIAPRESENCE ON ASIAN CARP AND NATIVE FISHES OF THEILLINOIS RIVER

Andrea K. Fritts1*, Alison P. Stodola2, Sarah A. Douglass2 & Rachel M. Vinsel2

1 Illinois Natural History Survey, Illinois River Biological Station, 704 N. Schrader Avenue, Havana,

IL 62644 USA2 Illinois Natural History Survey, 1816 S. Oak Street, Champaign, IL 61820 USA

ABSTRACT

Densities of introduced Asian carp (Silver Carp and Bighead Carp) in the Illinois River Basin areamong the highest in the world. Asian carp have been reported to serve as hosts for Sinanodontawoodiana in their native territories, but no research has been conducted on the potential for Silver orBighead Carp to host North American freshwater mussels. Our objectives were 1) to examine thepresence of glochidia on native and non-native fishes from the Illinois River Basin, 2) to determine anoptimal concentration and duration of potassium hydroxide (KOH) exposure for increasingtransparency of preserved fish gills to more effectively detect the presence of glochidia and parasites,and 3) identify parasite burdens. Fifteen fish species (12 native and 3 non-native) were collected fromthe Illinois River Basin during summer of 2014. Preserved fins and gills of native and non-native fisheswere examined for glochidia and parasite infections. We determined that a 20 min 5% KOH bath wasoptimal for increasing gill transparency. We recovered 242 glochidia from 5 native fish species: Bluegill,Largemouth Bass, Smallmouth Bass, Freshwater Drum, and Sauger. Based upon morphometric data,we were able to identify the glochidial larval stage of 5 groups of freshwater mussels: Group A-Lilliput,Group B- Threeridge, Group C- Deertoe or Fawnsfoot, Group D- Threehorn Wartyback, and Group E-Fragile Papershell. We did not locate glochidia on any of the non-native fish species. Future researchshould include the use of laboratory host trials to elucidate if Asian carp could serve as successful hostfishes for native mussels or if they are recruitment sinks, a possibility that could have a major impact onthe future stocks of currently imperiled freshwater mussels.

KEY WORDS - Unionoida, parasites, Hypophthalmichthys molitrix, Hypophthalmichthys nobilis, Cyprinuscarpio

INTRODUCTION

Freshwater mussels have experienced substantial declines in

their populations worldwide over the past century (Starrett

1971; Lydeard et al. 2004). However, there have been positive

developments in some locations where mussel populations have

been able to recolonize areas from which they had previously

been extirpated (Sietman et al. 2001). Unfortunately, the

success of some of these recolonizations may be undermined

by emerging threats. The introduction of Silver Carp (Hypo-phthalmichthys molitrix) and Bighead Carp (Hypophthalmich-thys nobilis), hereafter referred to collectively as Asian carp,

into North America has had a substantial negative impact on

native fish species, including declines in condition indices and

population size (Irons et al. 2007; Crimmins et al. 2015).

However, little research has been conducted to evaluate how

Asian carp are affecting other aquatic organisms.

The life cycle of freshwater mussels is complex and unique

among bivalves. Larval mussels (glochidia) are released by the

adult female and must attach to gills or fins of a suitable host

fish (Kat 1984; Barnhart et al. 2008). If it is an appropriate

host, glochidia remain attached to the fish for several days to

several weeks and metamorphose into juvenile mussels.

Juveniles are released from the host and fall to the benthic

substrates to continue their life cycle as free-living organisms.

Glochidia can also attach to non-suitable hosts but the fishes’

immune systems eventually reject the glochidia, which fall

from the fish and perish (Kat 1984).*Corresponding Author: andreakayfritts@gmail.com

Asian carp have been reported to serve as fish hosts to

freshwater mussels in their native range (Djajasasmita 1982;

Girardi and Ledoux 1989; Domagala et al. 2007). However, no

research has been conducted on the potential for Silver Carp

and Bighead Carp to host North American freshwater mussels

or to determine if they serve as recruitment ‘‘sinks.’’ Asian

carp densities in Illinois River Basin are among the highest in

the world (~2,500 per river km; Sass et al. 2010), and

glochidia may inadvertently attach to carp. If glochidia do not

metamorphose, Asian carp may be reproductive sinks that

prevent glochidia from attaching to suitable native host fishes.

We conducted a study to evaluate the potential effects of

invasive Asian carp on native freshwater mussels. Our primary

objective was to investigate the presence of glochidia on

native and non-native fishes throughout the Illinois River

Basin. Special emphasis was placed on examining three

species of carp (Silver, Bighead, and Common Carp (Cyprinuscarpio)), as these species may have the potential to intercept

glochidia and interfere with glochidial attachment to suitable

native fish hosts. Secondary objectives were to identify other

parasite burdens carried by native and non-native fishes, as

well as determining an optimal concentration and exposure

time for potassium hydroxide (KOH) solution to increase

clarity of preserved fish gills to more effectively locate and

identify encysted glochidia.

METHODS

Fishes were collected during May, June, and July of 2014,

which coincides with peak glochidial release for most

freshwater mussel species in the Illinois River Basin (Watters

et al. 2009). We targeted native fishes that coincidentally occur

in microhabitat with Asian carp, as well as natives that are

proven hosts for a variety of common mussels in the basin. We

also collected Common Carp, a non-native species that has

existed in the river since the 1800’s and has been reported to

be a marginal host for three North American freshwater mussel

species (Lefevre and Curtis 1910; Hove et al. 2011a; Hove et

al. 2014). Fishes were collected from the Upper Illinois River

(Dresden and Marseilles reaches), the Middle Illinois River

(La Grange Reach), as well as from several major tributaries to

the Illinois River (Kankakee, Spoon, Mackinaw, and Sanga-

mon rivers, and Salt Creek of the Sangamon).

Fishes were collected using a combination of pulsed-DC boat

electrofishing, hoop netting, and fyke netting and were then

returned to the laboratory for analysis. Smaller fish were

preserved whole in 95% ethanol, while gills and fins were

removed from larger fish (e.g., invasive carp) and were either

preserved in 95% EtOH or were frozen. All Animal Care and Use

protocols for fish collection, anesthetization, and euthanasia were

followed (University of Illinois IACUC #14023). Freezing gills

is not believed to influence detection of glochidia (Cunjak and

McGladdery 1991). Fish were identified to species and then gill

and fin tissues were examined for glochidia and other parasites

under a Leica S8 APO stereomicroscope (Leica Microsystems,

Wetzlar, Germany). When glochidia were found, they were

counted, photographed with a Leica DMC 2900 digital camera

(Leica Microsystems, Wetzlar, Germany), and measured for their

length (parallel to hinge), height (perpendicular to hinge), and

hinge length using Leica scale bar measurements in ImageJ

(version 1.46r). Glochidia were left in the gill tissue unless they

released from the tissue during processing. Efforts were made to

orient all of the glochidia onto a level plane prior to being

photographed, and only glochidia with suitable orientations were

used for measurements. To ensure accuracy of our measure-

ments, a subset of glochidia were measured by using three

different techniques (Leica scale bar measurements in ImageJ,

micrometer photograph measurements in ImageJ, and Leica

measurements using the Leica measurement software). Efforts

were made to identify the glochidia using morphological features

to the lowest taxonomic level possible. Based upon morphomet-

ric data and recent mussel community information from the

Illinois River, we used a bivariate plot and simple qualitative

overlap to determine the most-likely identity of each glochidium,

as our sample size was small (Kennedy and Haag 2005).

Morphometric data were derived from Waller (1987), Hoggarth

(1999), Williams et al. (2008) and references therein, Watters et

al. (2009), Hove et al. (2012), Hove et al. (2015) and M.C. Hove

(Macalester College, personal communication). When other

types of parasites were encountered, the parasite type and

infection intensity was also recorded for all of the native and non-

native fishes. We also documented the occurrence of telangiec-

tasia, a condition in which the blood vessels within the lamellae

of the gill filaments burst and blood pools in the lamellae tips,

causing the gills to have cyst-like structures along the gill

margins that superficially resemble parasitic infections. We

recorded the occurrence of these structures and compared the

probability of occurrence in native versus non-native fishes using

logistic regression (R Core Package 2015).

Potassium hydroxide study for gill clarification optimization

We completed an experiment to determine the most

effective processing treatment for detection of gill parasites.

Gill size was determined by measuring from dorsal to ventral

margin of an entire gill arch removed from the fish; small,

medium, and large size classes were approximately 10mm

(i.e., Spotfin Shiner, Cyprinella spiloptera), 10-50mm (i.e.,

Smallmouth Bass, Micropterus dolomieu), and greater than

50mm (i.e., Silver Carp), respectively. Three concentrations of

KOH were used: 2%, 5%, and 10% KOH. Transparency and

intactness of the gills were evaluated at six different time

points: 1, 5, 10, 20, 30, and 60 minutes. A black and white grid

(cell size of 10 3 10 mm) was used to record transparency by

determining when the transition line from black to white was

visible through the gill filament. Transparency was noted for

tips or edges of gill filament, mid-vein of filament, and whole

filament. Intactness was determined by picking up the gill and

recording if parts of it began to disintegrate.

23GLOCHIDIA PRESENCE ON ILLINOIS RIVER FISHES

RESULTS

Fishes were collected from 66 sites between 20 May 2014

and 22 July 2014 (Figure 1). We analyzed a total of 435 fishes

for this study, including 145 non-natives (92 Silver Carp, 3

Bighead Carp, and 50 Common Carp) and 290 native fishes

(12 species from 5 families) that are known to be host fish for

the larval stage of at least one freshwater mussel species

(Table 1). Of the total number of native fish collected, 10.7%

were infected with glochidia, and five native fish species were

infected. Infection rates among these species ranged from

12.5-100%. We recovered 242 glochidia from the 5 native fish

species: Bluegill (Lepomis macrochirus), Largemouth Bass

(Micropterus salmoides), Smallmouth Bass, Freshwater Drum

(Aplodinotus grunniens), and Sauger (Sander canadensis)

(Table 2). We identified the glochidial larval stage of 5 groups

of freshwater mussels: Group A-Lilliput (Toxolasma parvum),

Group B- Threeridge (Amblema plicata), Group C- Deertoe

(Truncilla truncata) or Fawnsfoot (Truncilla donaciformis),

Group D-Threehorn Wartyback (Obliquaria reflexa), and

Group E- Fragile Papershell (Leptodea fragilis) (Table 3,

Figure 2).

We recovered 9 types of non-glochidial parasites on 10 fish

species. These parasites included: anchor worms (Lernaeasp.), black grub (Neascus), white grub (Posthodiplostomumminimum), yellow grub (Clinostomum sp.), monogenean

trematodes (Dactylogyrus sp.), digenean trematode (Bolbo-phorus sp.), copepods, leeches, and nematodes. Telangiectasia

were documented in eight fish species, and this phenomenon

occurred more frequently in non-native fishes (Silver Carp and

Common Carp) (Figure 3, Table 4). Non-native fishes were

4.3 times more likely to have telangiectasia than native fishes

(95% confidence interval ¼ 2.9 to 7.9 times more likely).

Potassium hydroxide study for gill clarification optimization

The optimal concentration of KOH was determined to be a

5% KOH bath for 20 minutes. This concentration provided

maximum transparency without causing excessive gill tissue

deterioration. Smaller fishes and frozen Silver Carp gill

specimens reached a sufficient level of transparency in less

Figure 1: Fish collection sites on the Illinois River and its tributaries, Illinois,

USA.

Table 1. Native and non-native fishes collected in the Illinois River and its tributaries during May, June, and July of 2014.

Common name Scientific Name Tributaries Upper Illinois Middle Illinois

Gizzard Shad Dorosoma cepedianum 0 1 0

Red Shiner Cyprinella lutrensis 0 0 93

Spotfin Shiner Cyprinella spiloptera 5 31 0

Common Carp Cyprinus carpio 0 27 22

Silver Carp Hypophthalmichthys molitrix 53 0 39

Bighead Carp Hypophthalmichthys nobilis 1 0 2

Spottail Shiner Notropis hudsonius 0 0 9

Bluntnose Minnow Pimephales notatus 0 5 0

Bullhead Minnow Pimephales vigilax 0 9 31

Green Sunfish Lepomis cyanellus 0 1 0

Bluegill Lepomis macrochirus 0 29 28

Smallmouth Bass Micropterus dolomieu 0 4 0

Largemouth Bass Micropterus salmoides 0 14 5

Sauger Sander canadensis 0 1 0

Freshwater Drum Aplodinotus grunniens 0 0 24

24 FRITTS ET AL.

time, but the intactness of their gills was not negatively

affected after 20 minutes of KOH exposure.

DISCUSSION

We did not recover glochidia from any non-native fishes in

this study. This result was not entirely unexpected for a

number of reasons. First, if Asian carp were intercepting

glochidia but were not suitable hosts, it is likely that the

glochidia would slough from the carp within 1-4 days (Arey

1932a; Zale and Neves 1982a; Waller and Mitchell 1989). To

document attachment, we would need to have collected fishes

within a short period of time (i.e., ,4 days) after they

encountered the glochidia. Second, the Illinois River was in

flood stage for most of the summer during 2014. Any

glochidia that would have been present in the system,

particularly in the water column, would have been more

dilute than in a regular flow year. The flooding may have

contributed to us not finding glochidia on any of our pelagic

species (e.g., native cyprinids or Silver Carp). We also found

no evidence of natural infestation on the benthic dwelling

Common Carp. The gills of this species were rather difficult to

process; the densely packed gill filaments of the Common

Carp began to deteriorate in the KOH bath more quickly than

any other species. This phenomenon could have decreased our

ability to detect glochidia on Common Carp. Further, they are

only considered marginal hosts for three native mussel species

in Illinois that are considered host generalists: Rock

Pocketbook (Arcidens confragosus), Flutedshell (Lasmigonacostata), and Giant Floater (Pyganodon grandis) (Lefevre and

Curtis 1910; Hove et al. 2011a; Hove et al. 2014). Thus

glochidia would be subject to sloughing in a short time period

in this case as well.

The majority of the native species that were infested with

glochidia were collected in the Upper Illinois River, where

recent surveys have seen a considerable rebound in mussel

populations over the last decade (INHS Mollusk Collection;

http://wwx.inhs.illinois.edu/collections/mollusk; accessed 1

July 2015). This recovery may be a response to improved

water quality conditions brought forth by the Clean Water Act

of 1972, as an extensive mussel survey in 1966 revealed

extremely low mussel populations (Starrett 1971). Fifty-five

percent of the Bluegill and 50% of Largemouth Bass collected

from the Upper Illinois River were infected with glochidia.

The best fit mussel species for identified glochidia are from the

tribes Lampsilini and Amblemini. Lampsiline species typically

have high fecundity, shorter life spans, and are host specialists

that utilize only a single host fish species or a few species

within a genus or family. Fragile Papershell, Deertoe, and

Fawnsfoot are Freshwater Drum specialists and cannot

metamorphose on any other fish species. Lilliput likely use

Lepomis spp. and potentially darters as hosts (Mermilliod inFuller, 1978; Hove, 1995; Watters et al. 2005). Threehorn

Wartyback are known to metamorphose on cyprinids, but

marginal metamorphosis success on fishes from several

families has also been observed (Watters et al. 1998, B.R.

Table 2. Native fish species from the Upper and Middle Illinois River (ILR) with glochidia presence. N ¼ number of native fish collected.

Fish species Upper ILR Middle ILR Total N % infested No. glochidia Mean glochidia/fish

Bluegill 16 3 57 33% 178 9.4

Smallmouth Bass 1 0 4 25% 1 1.0

Largemouth Bass 7 0 19 37% 8 1.1

Sauger 1 0 1 100% 38 38.0

Freshwater Drum 0 3 24 13% 17 5.7

Table 3. Mean (61 SE) and ranges (in parentheses) for measurements of glochidia recovered from naturally infested fishes of the Upper and Middle Illinois River

(ILR). N¼ number of glochidia identified from each reach. Best fit species were assigned based upon established glochidial measurements from the literature and

knowledge of the current mussel assemblage present in the ILR. Fish species¼ species from which the glochidia were obtained.

Glochidia

group

Height

(lm)

Length

(lm)

Hinge

(lm)

N, Upper

ILR

N, Middle

ILR

Total

N

Best fit

species

Fish

species

A 176 6 1

(163-188)

158 6 1

(146-171)

87 6 1

(78-97)

52 0 52 Lilliput Bluegill, Sauger

B 208 6 1

(206-210)

187 6 1

(182-190)

134 6 1

(131-136)

3 2 5 Threeridge Bluegill, Largemouth

Bass, Freshwater Drum

C 54 6 1

(52-58)

57 6 3

(49-64)

36

(36-36)

0 7 7 Deertoe, Fawnsfoot Freshwater Drum

D 263 6 7

(251-281)

251 6 1

(243-259)

147 6 3

(139-154)

0 15 15 Threehorn Wartyback Bluegill

E 77 71 45 0 1 1 Fragile Papershell Freshwater Drum

25GLOCHIDIA PRESENCE ON ILLINOIS RIVER FISHES

Bosman, Texas Tech, personal communication). Amblemines

are typically longer-lived, slower to reach sexual maturity, and

can be host specialists or generalists (Watters et al. 2009). The

Threeridge is a host generalist that can utilize many fish

species, which is a factor that likely contributes to its

dominance among mussel fauna within the Illinois River

(Haag 2012).

We used a combination of morphological measurements and

current species distribution to determine the most likely species

assignment of attached glochidia. There were additional species

that did overlap morphometrically with our glochidia but were

eliminated based on known status of freshwater mussels in

Illinois. Specifically, Spike (Elliptio dilatata) and Spectaclecase

(Margaritifera monodonta) both have similar measurements as

our designated Groups B and C, respectively, but have not been

recovered alive from the Illinois since the early 20th century

(Starrett 1971, Sietman et al. 2001). However, there have been

recent discoveries of species considered extirpated, such as

Scaleshell (Leptodea leptodon) (Illinois Natural History Survey

Mollusk Collection; http://wwx.inhs.illinois.edu/collections/

mollusk; accessed 1 July 2015), thus we cannot completely

rule out the possibility that Spike or Spectaclecase may be

attached to native fishes in the Illinois River.

Glochidia size can vary widely within a species throughout

its range (Hove et al. 2011b, 2012, and M.C. Hove Macalester

College, personal communication). An optimal situation

would be to collect gravid females from the same waterbody

and compare known glochidia morphology with the unknown

glochidia from natural infestations. This was not an option in

this study due to the flooding/sampling conditions at the time

of fish collection; additionally, the availability of preserved

specimens with mature glochidia from the Illinois River is also

very limited, given the recent recolonization of this water

body. Future research involving naturally infested glochidia

should strive to include the collection of brooding females and

glochidia from the areas of interest to allow for more accurate

morphological comparisons.

In much of the literature with wild-caught fish, thousands

of fish were examined to elucidate glochidial infestation levels

(Zale and Neves 1982b; Neves and Widlak 1988; Weiss and

Figure 2. Bivariate plot of glochidia height and length, with assumed groups

labeled as A, B, C, D, or E. Published size ranges for known glochidia are

represented by labeled dotted lines.

Figure 3. The occurrence of telangiectasia in gill filaments on Silver Carp. Top

photo is of a freshly collected gill and the bottom photo is a gill that had been

preserved in 95% EtOH.

Table 4. Occurrence of telangiectasia on native and non-native fishes in the

Illinois River (ILR) and its tributaries. Total N¼ total number of fish collected

per species.

Fish species Tributaries Upper ILR Middle ILR Total N

Red Shiner – – 16 93

Common Carp – – 16 49

Silver Carp 7 – 39 92

Bullhead Minnow – – 1 40

Bluegill – – 3 57

Smallmouth Bass – 2 – 4

Largemouth Bass – – 1 19

Freshwater Drum – – 6 24

26 FRITTS ET AL.

Layzer 1995; Boyer et al. 2011). Thus, the fact that we found

glochidia on a large percentage of the relatively low numbers

of fishes examined during this study suggests that the Illinois

River mussel community is recovering. However, it also

means we would need to examine a substantial number of

Asian carp to truly rule out the possibility that carp are not

intercepting glochidia, since we only examined 145 carp

during this study.

Silver, Bighead, and Common Carp did not carry many

parasites, but Silver and Common Carp did have a substantially

higher occurrence of telangiectasia hemorrhages compared to

native fishes. This is a novel finding that is not well reported in

the literature. It is unknown as to why non-native fishes may

experience a higher degree of hemorrhages. There was also a

higher occurrence of this condition in main stem fishes compared

to tributaries, despite the fact that the fishes were collected with

the same protocols and electrofishing settings. Variation in

conductivity between the tributaries and the main stem may have

contributed to the pattern; conductivity in the tributaries was

generally lower than that in the main stem and pulsed DC-

electrofishing can be affected by different conductivities (M.W.

Fritts and R.M. Pendleton, Illinois Natural History Survey, C.

Morgeson, Eastern Illinois University, personal communica-

tion). This may contribute to the increased presence of

hemorrhages, but ultimately the cause of this phenomenon

remains unresolved and will need additional research.

Asian carp are voracious filter-feeding consumers that can

filter particles (e.g., plankton and algae) as small as 10-20 lm

(Jennings 1988; Smith 1989; Voros 1997). This feeding

behavior introduces the potential for Asian carp to be

consuming glochidia, an area of research that is currently

unstudied. We focused our laboratory efforts on the filament

structure of the gills, because this is the most likely location

for glochidial gill-attachment in native fishes (Arey 1932b).

However, the Asian carp could have been collecting glochidia

with the fused, sponge-like gill rakers and subsequently

ingesting the glochidia. Further studies should consider

examining the gill rakers in addition to the filaments. It is

unlikely that we would be able to detect glochidia in the

stomach contents, but use of advance dietary studies, such as

stable isotope analysis, may shed light on the potential for

Asian carp to be ingesting glochidia.

Future work should include conducting laboratory host trials

to evaluate the physiological ability of Asian carp to successfully

transform any of our native mussel species, focusing on both

host specialists and host generalists. This would be an efficient

method to determine whether the Asian carp could successfully

metamorphose glochidia to the juvenile life stage. If they are not

suitable hosts, we could determine the period before untrans-

formed glochidia are sloughed.

ACKNOWLEDGEMENTS

Funding for this research was provided by the University

of Illinois-National Great Rivers Research and Education

Center Challenge Grant. We thank A. Casper, K. Cummings,

J. DeBoer, M. Fritts, C. Morgeson, R. Pendleton, L. Solomon,

and J. Tiemann for assistance in the field and laboratory. We

thank J. Epifanio for his helpful input on this study and his

laboratory that provided some fish collection assistance.

Members of the U.S. Fish and Wildlife Service La Crosse

Fish Health Center provided assistance on identification of

assorted parasites. The manuscript was improved with reviews

by M. Fritts, M. Hove, and an anonymous reviewer. All

Animal Care and Use protocols for fish collection, anesthe-

tization, and euthanasia were followed (University of Illinois

IACUC #14023).

REFERENCES

Arey, L. B. 1932a. A microscopical study of glochidial immunity. Journal of

Morphology 53:201-221.

Arey, L. B. 1932b. The formation and structure of the glochidial cyst.

Biological Bulletin (Woods Hole) 62:212-221.

Barnhart, M. C., W. R. Haag, and W. N. Roston. 2008. Adaptations to host

infection and larval parasitism in Unionoida. Journal of the North

American Benthological Society 27:370-394.

Boyer, S. L., A. A Howe, N. W. Juergens, and M. C. Hove. 2011. A DNA-

barcoding approach to identifying juvenile freshwater mussels (Bivalvia:

Unionidae) recovered from naturally infested fishes. Journal of the North

American Benthological Society 30:182-194.

Crimmins, S. M., P. Boma, and W. E. Thogmartin. 2015. Projected risk of

population declines for native fish species in the upper Mississippi River.

River Research and Applications 31:135-142.

Cunjak, R. A., and S. E. McGladdery. 1991. The parasite-host relationship of

glochidia (Mollusca: Margaritiferidae) on the gills of young-of-the-year

Atlantic salmon (Salmo salar). Canadian Journal of Zoology 69:353-358.

Djajasasmita, M. 1982. The occurrence of Anodonta woodiana Lea, 1837 in

Indonesia (Pelecypoda: Unionidae). Veliger 25:175.

Domagala, J., A. M. Labecka, B. Migdalska, and M. Pilecka-Rapacz. 2007.

Colonization of the channels of Miedzyodrze (north-western Poland) by

Sinanodonta woodiana (Lea, 1834) (Bivalvia: Unionidae). Polish Journal

of Natural Sciences 22:679-690.

Fuller, S. L. H. 1978. Fresh water mussels (Mollusca: Bivalvia: Unionidae) of

the upper Mississippi River: Observations at selected sites within the 9

foot channel navigation project on behalf of the U.S. Army Corps of

Engineers. Final Report to the U.S. Army Corp of Engineers, No. 78-33.

Girardi, H., and J. C. Ledoux. 1989. Presence d’Anodonta woodiana (Lea,

1834) en France (Mollusques, Lamellibranches, Unionidae). Bulletin

Mensuel de la Societe Linneenne de Lyon 58:286-290.

Haag, W. R. 2012. North American freshwater mussels: natural history,

ecology, and conservation. Cambridge University Press, Cambridge, U.K.

Hoggarth, M. A. 1999. Descriptions of some of the glochidia of the Unionidae

(Mollusca: Bivalvia). Malacologia 41:1–118.

Hove, M. C. 2005. Suitable fish hosts of the lilliput, Toxolasma parvus.

Triannual Annual Report 8:9.

Hove, M. C., B. E. Sietman, J. E. Bakelaar, J. A. Bury, D. J. Heath, V. E. Pepi,

J. E. Kurth, J. M. Davis, D. J. Hornbach, and A. R. Kapuscinski. 2011a.

Early life history and distribution of pistolgrip (Tritogonia verrucosa

(Rafinesque, 1820)) in Minnesota and Wisconsin. American Midland

Naturalist 165:338-354.

Hove, M., A. Fulton, K. Wolf, B. Sietman, D. Hornbach, S. Boyer, and N.

Ward. 2011b. Additional suitable hosts identified for Rock Pocketbook

(Arcidens confragosus). Ellipsaria 13:6-7.

27GLOCHIDIA PRESENCE ON ILLINOIS RIVER FISHES

Hove, M. C., M. T. Steingraeber, T. J. Newton, D. J. Heath, C. L. Nelson, J. A.

Bury, J. E. Kurth, M. R. Bartsch, W. S. Thorpe, M. R. McGill, and D. J.

Hornbach. 2012. Early life history of the winged mapleleaf mussel

(Quadrula fragosa). American Malacological Bulletin 30(1):47-57.

Hove, M., B. Davis, E. Wanner, P. Leonard, G. Van Susteren, B. Sietman, S.

Bump, S. Marr, K. Murphy, M. Berg, A. Handy, A. Thoreen, K. Rod, M.

Hanson, R. Pochman, S. Nelson, and W. LaMere. 2014. Laboratories

show Lasmigona costata metamorphose on several fish species. Ellipsaria

16:21-23.

Hove, M. C., B. E. Sietman, M. S. Berg, E. C. Frost, K. Wolf, T. R. Brady, S.

L. Boyer, and D. J. Hornbach. 2015. Early life history of the sheepnose

(Plethobasus cyphyus) (Mollusca: Bivalvia: Unionoida). Journal of

Natural History. http://dx.doi.org/10.1080/00222933.2015.1083059

Irons, K. S., G. G. Sass, M. A. McClelland, and J. D. Stafford. 2007. Reduced

condition factor of two native fish species coincident with invasion of

non-native Asian carps in the Illinois River, U.S.A. Is this evidence for

competition and reduced fitness? Journal of Fish Biology 71:258-273.

Jennings, D. P. 1988. Bighead carp (Hypophthalmichthys nobilis): a biological

synopsis. U.S. Fish Wildlife Service, Biology Report 88:1–35.

Kat, P. W. 1984. Parasitism and the Unionacea (Bivalvia). Biological Reviews

of the Cambridge Philosophical Society 59:189-207.

Kennedy, T. B., and W. R. Haag. 2005. Using morphometrics to identify

glochidia from a diverse freshwater mussel community. Journal of the

North American Benthological Society 24:880–889.

Lefevre, G., and W. C. Curtis. 1910. Reproduction and parasitism in the

Unionidae. Journal of Experimental Zoology 9:79-116.

Lydeard, C., R. H. Cowie, W. F. Ponder, A. E. Bogan, P. Bouchet, S. A. Clark,

K. S. Cummings, T. J. Frest, O. Gargominy, D. G. Herbert, R. Hershler,

K. E. Perez, B. Roth, M. Seddon, E. E. Strong, and F. G. Thompson.

2004. The global decline of nonmarine mollusks. Bioscience 54:321–330.

Neves, R. J., and J. C. Widlak. 1988. Occurrence of glochidia in stream drift

and on fishes of the Upper North Fork Holston River, Virginia. American

Midland Naturalist 119:111-120.

R Core Team. 2015. R: A language and environment for statistical computing.

R Foundation for Statistical Computing, Vienna, Austria. URL https://

www.R-project.org/.

Sass, G. G., T. R. Cook, K. S. Irons, M. A. McClelland, N. N. Michaels, T. M.

O’Hara, and M. R. Stroub. 2010. A mark-recapture population estimate

for invasive silver carp (Hypophthalmichthys molitrix) in the La Grange

Reach, Illinois River. Biological Invasions 12:433-436.

Sietman, B. E., S. D. Whitney, D. E. Kelner, K. D. Blodgett, and H. L. Dunn. 2001.

Post-extirpation recovery of the freshwater mussel (Bivalvia: Unionidae)

fauna in the upper Illinois River. Journal of Freshwater Ecology 16:273-281.

Smith, D. W. 1989. The feeding selectivity of silver carp, Hypophthalmichthys

molitrix Val. Journal of Fish Biology 34:819–828.

Starrett, W. C. 1971. A survey of the mussels (Unionacea) of the Illinois River: a

polluted stream. Illinois Natural History Survey Bulletin 30:267-403.

Voros, L. 1997. Size-selective filtration and taxon-specific digestion of

plankton algae by silver carp (Hypophthalmichthys molitrix Val.).

Hydrobiologia 342:223–228.

Waller, D. R. 1987. Studies on Lampsilis mussels of the Upper Mississippi

River. Doctoral Dissertation. Iowa State University, Ames, Iowa.

Waller, D. L., and L. G. Mitchell. 1989. Gill tissue reactions in walleye

Stizostedion vitreum and common carp Cyprinus carpio to glochidia of

the freshwater mussel Lampsilis radiata siliquoidea. Diseases of Aquatic

Organisms 6:81-87.

Watters, G. T., S. H. O’Dee, S. Chordas, and J. Reiger. 1998. Potential hosts

for Lampsilis reeviana brevicula, Obliquaria reflexa. Triannual Unionid

Report 16:21-22.

Watters, G. T., T. Menker, S. Thomas, and K. Kuehnl. 2005. Host

identifications or confirmations. Ellipsaria 7(2):11-12.

Watters, G. T., M. A. Hoggarth, and D. H. Stansbery. 2009. The freshwater

mussels of Ohio. Ohio State University Press, Columbus, Ohio.

Weiss, J. L., and J. B Layzer. 1995. Infestations of glochidia on fishes in the

Barren River, Kentucky. American Malacological Bulletin 11:153-159.

Williams, J. D., A. E. Bogan, and J. T. Garner. 2008. Freshwater mussels of

Alabama and the Mobile Basin in Georgia, Mississippi, and Tennessee.

The University of Alabama Press, Tuscaloosa, Alabama.

Zale, A. V., and R. J. Neves. 1982a. Fish hosts of four species of lampsiline

mussels (Mollusca: Unionidae) in Big Moccasin Creek, Virginia.

Canadian Journal of Zoology 60:2535-2542.

Zale, A. V., and R. J. Neves. 1982b. Identification of a host fish for

Alasmidonta minor (Mollusca: Unionidae). American Midland Naturalist

107:386-388.

28 FRITTS ET AL.

Freshwater Mollusk Biology and Conservation 19:29–35, 2016

� Freshwater Mollusk Conservation Society 2016

ARTICLE

TOXICITY OF SODIUM DODECYL SULFATE TOFEDERALLY THREATENED AND PETITIONEDFRESHWATER MOLLUSK SPECIES

Kesley J. Gibson1,3, Jonathan M. Miller1, Paul D. Johnson2 & Paul M. Stewart1*

1 Troy University, Department of Biological and Environmental Sciences, 223 MSCX, Troy, AL

36082 USA2 Alabama Aquatic Biodiversity Center, 2200 Highway 175, Marion, AL, 36756 USA

ABSTRACT

Anthropogenically caused physical and chemical habitat degradation, including water pollution,have caused dramatic declines in freshwater mollusk populations. Sodium dodecyl sulfate (SDS), asurfactant with no USEPA Water Quality Criteria (WQC), is commonly used in industrial applications,household cleaners, personal hygiene products, and herbicides. In aquatic habitats, previous SDSstudies have associated deformities and death to mollusks found in these systems. The objective of thisstudy was to determine EC50 values for two freshwater juvenile unionids (Villosa nebulosa and Hamiotaperovalis) and two freshwater caenogastropods (Leptoxis ampla and Somatogyrus sp.) endemic to theMobile River Basin, USA, to SDS. Using the Trimmed Spearman-Karber method, EC50 values werecalculated. Results found that EC50 values were: V. nebulosa¼14,469 lg/L (95% CI: 13,436 – 15,581 lg/L), H. perovalis ¼ 6,102 lg/L (95% CI: 4,727 – 7,876 lg/L), Somatogyrus sp. ¼ 1,986 lg/L (95% CI:1,453 – 2,715 lg/L), and L. ampla¼ 26 lg/L (95% CI: 6 – 112 lg/L). Freshwater gastropods were moresensitive to SDS than freshwater unionids. Leptoxis ampla was the most sensitive species tested and hadsuch a low EC50 value that more protective regional criteria may be required. Therefore, futureresearch should include additional testing on mollusk species, particularly regionally isolated speciesthat may display increased sensitivity.

KEY WORDS - SDS, threatened, mollusk, Mobile River Basin, Water Quality Criteria

INTRODUCTION

In North America, freshwater mollusks are the most

imperiled aquatic fauna with 74% of 703 identified gastropods

imperiled, followed by unionids with 72% of 298 identified

species imperiled (Wilcove and Master 2008; Johnson et al.

2013). Many freshwater mollusk species are highly endemic,

particularly in the Mobile River Basin, USA, which includes

139 endemic freshwater mollusk taxa (34 unionids and 105

gastropods) (Neves et al. 1997; Williams et al. 2008; O Foighil

et al. 2011). Stenotypic species are often underrepresented in

traditional toxicity testing that normally utilize broad ranging

species, usually distributed across multiple drainage basins (O

Foighil et al. 2011; Wang et al. 2010; Wang et al. 2011).

Declines in freshwater gastropod and unionid populations

are attributed to increases in the human population, leading to

alteration or destruction of habitat both physically and

chemically (Villella et al. 2004; Johnson et al. 2013). Pollution

is ranked as the second leading cause of stream impairment

(USEPA 2009), following physical habitat alteration (Neves et

al. 1997). Toxicity testing is important in protecting organisms

by providing information on specific pollutant effects, such as

reduced survival and growth or inhibited biological processes

on a particular life stage (American Society for Testing and

Materials (ASTM) 2013). Using data from these tests, criteria

inclusive of imperiled organisms can be established to help

protect remaining populations.

*Corresponding Author: mstewart@troy.edu3Current Address: Texas A&M University- Corpus Christi, Harte

Research Institute, 6300 Ocean Drive, Corpus Christi, TX, 78412 USA

Sodium dodecyl sulfate (SDS), a surfactant, is the most

widely used synthetic organic chemical found in detergents,

shampoos, cosmetics, household cleaners, herbicides, and

dispersants used in oil-spill cleanups (Cowan-Ellsberry et al.

2014). Sodium dodecyl sulfate is an alkylsulfate with sodium

as the counter ion with a chain length of 12 carbons (Cowan-

Ellsberry et al. 2014). Sodium lauryl sulfate (SLS) and SDS

are often used synonymously in reporting of product

ingredients (Singer and Tjeerdema 1993; NIH, 2014).

Concentrations .67% SLS (active ingredients) can be found

in household products, dispersants, and herbicides (Lewis

1991; Singer and Tjeerdema 1993; Kegley et al. 2014).

Sodium dodecyl sulfate is also used in the cleanup of

polycyclic aromatic hydrocarbons (e.g., oil and gas products).

The major exposure route for SDS to aquatic environments is

through contaminated waters, sediments, or soils, which

threatens drinking water supplies or organisms living in these

environments (Singer and Tjeerdema 1993). Contamination of

groundwater by surfactants is caused primarily by leaching

from industrial and municipal sewage systems, but can also be

introduced to the environment by domestic and industrial

effluents from discarded cleaning products (Singer and

Tjeerdema 1993; Chaturvedi and Kumar 2010). In the United

States, per capita detergent consumption is about 10 kg/year

(Rebello et al. 2014), but consumption declined 3.9% per year

during 2008 – 2013 (Cowan-Ellsberry et al. 2014). However,

in 2008, 76% of alkylsulfate consumed in North America were

found in household laundry detergents (59%) and personal

care products (17%) (Cowan-Ellsberry et al. 2014).

Sodium dodecyl sulfate is not currently monitored in water

systems or listed as a ground water contaminant (Kegley et al.

2014). Other surfactants with similar uses are monitored

(reviewed by Rebello et al. 2014 and Singer and Tjeerdema

1993). In the United Kingdom, surfactant concentrations in

surface waters have been recorded as high as 416 lg/L (Fox et

al. 2000), while sewage effluents have had concentrations

documented up to 1,090 lg/L (Holt et al. 1989). Treated

sludge has been found to have concentrations of linear

alkylbenzene sulfonate as high as 30,200,000 lg/kg (dry

weight) (Berna et al. 1989). All monitored concentrations for

sulfates exceed the predicted no-effect concentration value

(250 lg/L) for surfactants by van de Plassche et al. (1999). In

Massachusetts, the Town River had reported concentrations

between 40 lg/L and 590 lg/L (Lewis and Wee 1983), while

other major rivers in the United States had reported surfactant

concentrations that ranged from 10 lg/L to 3,300 lg/L (A.D.

Little Co. 1981) or 10 lg/L to 40 lg/L (Hennes and Rapaport

1989).

Sodium dodecyl sulfate was formally classified as

‘environmental friendly’ based on its readily biodegradable

and low bioaccumulation properties, meaning it does not

persist long in the environment (Belanger et al. 2004).

However, some studies have suggested that SDS can be lethal

in acute exposures (e.g., 19,040 lg/L for Utterbackiaimbecilis, Keller 1993; summarized in Singer and Tjeerdema

1993; summarized in Kegley et al. 2014; Table 2). Because of

its fast acting, nonselective, and consistent toxicity, SDS is

commonly used as a reference toxicant in toxicity tests

(USEPA 2002). Developmental abnormalities in Illyanassaobsoleta embryos, such as incomplete or inhibited formation

of lobe-dependent structures (e.g., foot, operculum, and eyes)

of gastropods have been attributed to SDS exposure

(treatments ranging from 10,000 – 30,000 lg/L) (Render

1990). Tarazona and Nunez (1987) reported that SDS

exposure significantly decreased shell weights in lymnaeid

gastropods and impeded normal shell deposition (EC50¼ 540

lg/L for Lymnaea vulgaris and 610 lg/L for Physaheterostropha). When exposed to SDS (EC50 ¼ 31,400 lg/

L), Corbicula fluminea displayed avoidance behaviors and gill

damage which decreased oxygen consumption and reduced

siphoning activity (Graney and Giesy 1988).

Previous studies suggest early life stages of unionids are

more sensitive than later life stages or other commonly used

aquatic test organisms (Keller et al. 2007; Augspurger 2013).

Until recently, freshwater unionid toxicity tests were not

included in establishing Water Quality Criteria (WQC) due to

limited information available (e.g., life cycle, host fish,

sensitivity, populations), and the inability to culture them in

sufficient numbers to support testing needs (Keller et al. 2007).

Table 1. Data from acute toxicity trials using SDS on four mollusk species,

including toxicant concentrations, number of dead organisms, and number of

organisms exposed.

Species

Concentration

(lg/L) Dead

Number

exposed

Villosa nebulosa Control 3 10

5,000 1 30

10,000 1 30

15,000 16 30

20,000 28 30

30,000 30 30

Hamiota perovalis Control 0 10

5,000 11 30

10,000 16 30

15,000 17 30

20,000 28 30

30,000 30 30

Leptoxis ampla Control 0 10

1 3 30

10 11 30

100 21 30

1,000 23 30

10,000 18 30

Somatogyrus sp. Control 0 10

1,000 7 30

3,000 20 30

10,000 30 30

30,000 30 30

100,000 30 30

30 GIBSON ET AL.

Similarly, caenogastropods (respire using a gill or ctenidia) are

considered among the most sensitive aquatic organisms to

contaminants (Besser et al. 2009), but are rarely used for

toxicity testing due to slow growth and low reproductive rates,

which make them difficult to culture or test in the laboratory

(Besser et al. 2009). New propagation and rearing techniques

have been recently developed that allow sufficient numbers of

organisms to support formal toxicity testing (Barnhart 2006).

The goal of the current study was to evaluate acute SDS

exposure to four Mobile River Basin endemic mollusks, two

freshwater juvenile unionids and gastropods. These data may

eventually contribute to the development of a specific WQC

for SDS for freshwater mollusks.

METHODS

Two lotic freshwater unionids (Hamiota perovalis and

Villosa nebulosa) and two lotic freshwater caenogastropods

(Leptoxis ampla and Somatogyrus sp.) endemic to the Mobile

River Basin were used. Hamiota perovalis (Orangenacre

Mucket) and Leptoxis ampla (Round Rocksnail) were

federally listed as threatened under the Endangered Species

Act (ESA) (USFWS 1993, 1998). Villosa nebulosa (Alabama

Rainbow) has been formally petitioned for federal protection

under the ESA (Center for Biological Diversity 2010). The

specific taxonomic position of Somatogyrus sp., Cahaba

Pebblesnail, is unclear (E.E. Strong and P.D. Johnson,

Alabama Aquatic Biodiversity Center (AABC), personal

observation).

Unionids were propagated by the AABC, Marion,

Alabama, using host-fish infections and standard culturing

methods (Barnhart 2006). Villosa nebulosa adults (n¼6) were

collected from South Fork Terrapin Creek in Celburne County,

Alabama while H. perovalis adults (n¼4) were collected from

Rush Creek in Winston County, Alabama. Both species used

Largemouth Bass (Micropterus salmoides) for transformation.

Unionids were fed a diet of Nanochloropsis, Shellfish diet

(Reed Mariculture) at a concentration of ~50K cells/mL.

For gastropods, Leptoxis ampla was propagated by AABC,

while Somatogyrus sp. was collected from the Cahaba River

(Latitude: 328 57.5770 N, Longitude: 878 08.4410 W). The

AABC commonly uses this location for reintroductions,

restocking, and translocations of threatened and endangered

mollusk species. Juvenile unionids were 30 – 60 days post-

transformation, and gastropods were 5 – 8 months post hatch.

Somatogyrus sp. is considered to be an annual species with

juveniles hatching in spring. Adults die soon after the

reproductive season is concluded (Johnson et al. 2013). Test

organisms were kept in an aquarium with dilution water

prepared following ASTM (2007) guidelines and used in

testing within 14 days of arrival, so feeding was not necessary

(ASTM 2013). Since it can absorb contaminants (Newton and

Bartsch 2007), sediment was not used during toxicity testing,

which reduced the likelihood of organisms being exposed to

contaminants that may be present in sediment or uncontrolled

chemical reactions occurring within the sediments.

Experimental Conditions

Static renewal toxicity tests for both classes of organisms

were completed following ASTM Standard Guide for

Conducting Laboratory Toxicity Tests with Freshwater

Mussels (E2455-06) (ASTM 2013). Dilution water recipe

additions included sodium bicarbonate (NaHCO3), calcium

sulfate (CaSO4 �H2O), magnesium sulfate (MgSO4), and

potassium chloride (KCl) following ASTM Standard Guide

for Conducting Acute Toxicity Tests on Test Materials with

Fishes, Macroinvertebrates, and Amphibians (E729-96)

(ASTM 2007). The mean physiochemical variables of the

dilution water were as follows: pH - 7.3 (7.1 – 7.4), hardness -

Table 2. Median effective concentrations (EC50) for 96 h acute toxicity tests of sulfate surfactants on mollusk species.

Mollusk species Common name Age Hardness (mg CaCO3/L) 96 h EC50 (lg/L) References

Bivalves

Corbicula fluminea Asiatic Clam NR NR 31,400 Graney and Gisey 1988

Utterbackia imbecillis Paper Pondshell Juveniles NR 19,040a Keller 1993

Hamiota perovalis Orangenacre Mucket Juveniles 43 6,102 Current Study

Villosa nebulosa Alabama Rainbow Juveniles 43 14,469 Current Study

Gastropods

Physa heterostropha* Pewter Physa NR NR 34,161 Patrick et al. 1968

Lymnaea peregra* Wandering Pond Snail NR NR 15 Misra et al. 1984

Lymnaea vulgaris* Great Pond Snail Juveniles 35.6 540 Tarazona and Nunez 1987

Lymnaea vulgaris* Great Pond Snail Juveniles 35.6 610 Tarazona and Nunez 1987

Leptoxis ampla Round Rocksnail Juveniles 43 26 Current Study

Somatogyrus sp. Cahaba Pebblesnail Adults 43 1,986 Current Study

NR ¼ not reporteda ¼ 48 h LC50 value

* ¼ pulmonate gastropod

31TOXICITY OF SODIUM DODECYL SULFATE TO FRESHWATER MOLLUSKS

41 (40 – 42) mg CaCO3/L, alkalinity - 33.5 (33 – 34) mg

CaCO3/L, and conductivity - 168 (130 – 189) lX/cm.

Dissolved oxygen saturation measured �90%, temperature

was kept at 25 6 1 8C, and ambient light from overhead

fluorescent laboratory lights was used with a photoperiod of

16L: 8D. Test organisms were acclimated in the dilution water

for at least 24 hours before trials by adjusting the temperature

no more than 3 8C/h until reaching 25 8C (Wang et al. 2007;

ASTM 2013). Ten individuals were used in triplicate per

concentration (n ¼ 30). Chambers (600 mL Pyrext beakers)

were filled with 300 mL of either dilution water (controls) or

toxicant solution and were changed at 48 h (Table 1). Stock

SDS toxicant solutions (Laboratory grade, Lot # AD-14008-

56, Carolina Biological Supply) analyses were not conducted

to quantify concentrations in toxicant exposure; therefore,

EC50 values are based on nominal concentrations.

Endpoint determinations were completed after the 96 h

tests, unionids were placed under a microscope to view

heartbeat or foot movement. If no movement was observed

after five-minutes, the unionid was classified as dead.

Gastropods were classified as dead if no movement was

detected within a five-minute observation period (Archambault

et al. 2015) or after the ‘‘tickle’’ test, performed by touching

the organisms with a soft pick to provoke a stimulus response.

An eyelash stick was used to prevent any excess pressure

being placed on the foot and observing a false reaction. Non-

decaying individuals were placed in fresh dilution water for

thirty minutes and rechecked for survival. Control survivor-

ship had to exceed 90% at the end of each trial for results to be

acceptable (ASTM 2013) (Table 1).

Data Analysis

The 96 hour EC50 values for SDS were determined using

ToxStatt 3.5 from West, Inc. (downloaded from https://www.

msu.edu/course/zol/868/) using the Trimmed Spearman-Karb-

er method (TSK). The EC50 values of test organisms were then

compared to EC50 values of other species in the published

literature.

RESULTS

In accordance with ASTM (2013) protocols, control

survivability was �90% for each unionid trial and 100% for

each gastropod trial. Gastropods were active and scaling the

beaker walls within control test chambers. The EC50 value for

Hamiota perovalis was 6,102 lg/L (95% CI: 4,727 – 7,876

lg/L), and Villosa nebulosa was 14,469 lg/L (95% CI: 13,436

– 15,581 lg/L), more than double the EC50 value of H.perovalis (Table 2). In all treatments containing SDS, juvenile

unionids of both species purged a mucus-like substance that

coated the entire organism. While little foot movement was

observed, a heartbeat was always detected in live unionids.

However, in high concentrations of SDS, no soft tissue was

observed in the shells at the end of the 96 h acute toxicity tests.

Gastropods tested in the current study were more sensitive

to SDS than unionid species evaluated. Leptoxis ampla had an

EC50 value of 26 lg/L (95% CI: 6 – 112 lg/L), which was the

lowest EC50 value calculated in this study. Somatogyrus sp.

had an EC50 value of 1,986 lg/L (95% CI: 1,453 – 2,715 lg/

L) (Table 2). Similar to unionids, soft tissues dissolved or

completely separated from the shell in the highest concentra-

tions, and most dead gastropods had begun to decompose so

death was easily determined. Living gastropods from lower

concentrations appeared to begin normal activity once

transferred to water lacking SDS. Movement was observed

without ‘‘tickling’’ in most low concentrations.

DISCUSSION

Sodium dodecyl sulfate is commonly found in high

concentrations in detergents and household cleaners and

contaminates drinking water and aquatic ecosystems (Cow-

an-Ellsberry et al. 2014); however, it has no WQC. Keller

(1993) reported a 48 h LC50 value of 19,040 lg/L for juvenile

Utterbackia imbecillis, a species classified as having a stable

conservation status (Williams et al. 2008). Both juvenile

unionid species exposed to SDS in the current study had 96 h

EC50 values (V. nebulosa: 14,469 lg/L (federally petitioned);

H. perovalis: 6,102 lg/L (federally threatened)) below the

value reported by Keller (1993). In a related study, Graney and

Giesy (1988) reported a 96 h LC50 value of 31,400 lg/L for

SDS using Corbicula fluminea (Asiatic clam), which is 2x and

5x higher than the EC50 values determined for V. nebulosa and

Hamiota perovalis, respectively, in the current study.

The gastropod species used in the current study, L. amplaand Somatogyrus sp., were generally more sensitive to SDS as

compared to other published studies, but Tarazona and Nunez

(1987) reported a 96 h LC50 value of 540 lg/L for SLS using

Lymnaea vulgaris, a pulmonate gastropod. This reported value

was higher than the 96 h EC50 value for Leptoxis amplareported in the current study (26 lg/L), but Somatogyrus sp.

had a higher 96 h EC50 value at 1,986 lg/L, suggesting it may

be more tolerant than Lymnaea vulgaris. Patrick et al. (1968)

reported a LC50 value of 34,161 lg/L for alkylbenzene

sulfonate using the pulmonate gastropod Physa heterostropha,

which was a greater concentration than any EC50 value

reported in this study. Misra et al. (1984) reported an EC50

value of 15 lg/L (endpoint: calcium uptake) for alkylbenzene

sulfonate using Lymnaea peregra, which was similar to the

EC50 value for Leptoxis ampla. No other published EC50 or

LC50 values were close to the EC50 value of Leptoxis ampla,

suggesting that this highly stenotypic species (Cahaba River

Basin endemic) could be one of the most sensitive aquatic

species to SDS tested to date. These reported values suggest

that caenogastropods display increased sensitivity over

pulmonate gastropods to SDS contamination.

Fish have more frequently been subjected to SDS acute

toxicity testing than freshwater mollusks and tend to have

slightly higher LC50 values than mollusks (Table 3). However,

32 GIBSON ET AL.

invertebrates tend to be more sensitive than fish to SDS, and

few toxicity studies have focused on freshwater unionids using

SDS. The cladoceran Daphnia magna had 48 h LC50 value of

10,300 lg/L (Keller 1993), while Daphnia pulex had 48 h

LC50 values ranging between 1,400 lg/L and 15,200 lg/L

(Lewis and Weber 1985; Singer and Tjeerdema 1993) (Table

3). These lower values compared to the EC50 values of

Somatogyrus sp. reported in the current study suggest that D.pulex and Somatogyrus sp. may have similar SDS sensitivity.

Few toxicity studies have been conducted on caenogastropods

(Besser et al. 2009; Johnson et al. 2013), and previous

investigations with surfactants have used pulmonate gastro-

pods (e.g., Patrick et al. 1968; Misra et al. 1984; Tarazona and

Nunez 1987).

Other surfactants have been studied more extensively on

freshwater mollusks. Ostroumov and Widdows (2006) exam-

ined surfactants hindering filter feeding for unionids by

reporting a drop in clearance rates after a 10 minute exposure.

Bringolf et al. (2007b) examined the components of Roundupt

using Lampsilis siliquoidea and found that MON 0818, the

polyethoxylated tallow amine (POEA) surfactant blend that

helps the active ingredients penetrate the waxy coverings of

plant leaves, was the most toxic component of that herbicide.

In a similar study, Bringolf et al. (2007a) further examined

MON 0818 and reported that unionids during their earlier life

stages (i.e., glochidia and juvenile stages) are among the most

sensitive organisms tested.

The EC50 values of the current study were lower than most

reported in the literature (Tables 2 – 3) for SDS, suggesting

that some freshwater mollusks may be among the most

sensitive aquatic species tested to date. The majority of

mollusk species previously tested have broad geographical

ranges and occur across multiple basins and are, therefore,

probably adapted to a broad range of physical and chemical

water quality. In contrast, many federally listed freshwater

mollusks have distributions limited to a distinct, regional

drainage, such as the Mobile River Basin species utilized in

the current study. These regionalized species would be adapted

to specific regional physical and chemical parameters.

Freshwater unionids and caenogastropods in the current study

were found to be sensitive to SDS, particularly the Cahaba

River Basin endemic, Leptoxis ampla had the lowest LC50

value recorded to date. Threatened species, such as L. amplaor H. perovalis, often demonstrate increased sensitivity to

environmental changes than other broad ranging species. The

stenotypic biology of small range endemic mollusks with

unique generic placement (e.g., Hamiota) may represent

increased sensitivity to a variety of toxicants (Gibson 2015)

than wider ranging species adapted to a broad range of normal

water quality variables (e.g., Lampsilis siliquoidea).

Toxicity tests are important tools that provide information

for risk assessment of chemicals and are used when

determining USEPA WQC. This study is one of the few

testing toxicity of SDS using Mobile River Basin freshwater

mollusks, specifically using federally threatened or petitioned

species. Many mollusks have a narrow endemic range which

Table 3. Median effective concentrations (EC50) for 96 h acute toxicity tests of sulfate surfactants on macroinvertebrate and fish species.

Species name Common Name Hardness (mg CaCO3/L) 96 h EC50 (lg/L) References

Invertebrates

Daphnia magna Cladoceran NR 10,300 Keller 1993

Daphnia magna Cladoceran NR 5,400 – 15,000a Lewis and Weber 1985

Daphnia pulex Cladoceran NR 1,400 – 15,200a Lewis and Weber 1985

Fish

Danio rerio Zebrafish NR 7970 Fogels and Sprague 1977

Danio rerio Zebrafish NR 8810a Fogels and Sprague 1977

Danio rerio Zebrafish NR 9,900 – 20,100 Newsome 1982

Cichlasoma nigrofaciatum Convict Cichlid NR 16,100 – 30,000 Newsome 1982

Ctenopharyngodon idella Grass Carp NR 7,700 Susmi et al. 2010

Cynopoecilus melanotaenia Killfish NR 14,900 Arenzon et al. 2003

Cyprinus carpio Common Carp NR 1,310 Verma et al. 1981

Jordanella floridae Flagfish NR 8,100 Fogels and Sprague 1977

Macrones vittatus Asian Striped Catfish NR 1,390 Verma et al. 1978

Menidia beryllina Inland silverside NR 9,500 Hemmer et al. 2010

Oncorhynchus mykiss Rainbow Trout NR 4,620 Fogels and Sprague 1977

Piaractus brachypomus Red Pacu 24 11,290 Reategui-Zirena et al. 2013

Pimephales promelas Fathead Minnow NR 6,600 Conway et al. 1983

Pimephales promelas Fathead Minnow NR 10,000 – 22,500 Newsome 1982

Pimephales promelas Fathead Minnow NR 8,600 USEPA 2002

Poecilia reticulartus Guppy NR 13,500 – 18,300 Newsome 1982

NR ¼ not reporteda ¼ 48 h LC50 value

33TOXICITY OF SODIUM DODECYL SULFATE TO FRESHWATER MOLLUSKS

may increase sensitivity to water quality as compared to the

broader ranging species. Further testing is urged for regional

mollusk species, and especially determination of WQC for

SDS, which currently does not exist. Criteria may need to be

established using a suite of organisms from various river

basins to include stenotypic species (e.g., Cahaba River Basin

endemics) that may display increased sensitivity to SDS.

ACKNOWLEDGEMENTS

The authors wish to thank Dr. Chris Barnhart (Missouri

State University), Dr. Ning Wang (USGS), and Jennifer

Archambault (North Carolina State University) for their

assistance with this project. This manuscript benefited greatly

from input from two anonymous reviewers. This research was

done under FWS permit #TE130300-4. Funding was provided

by the ALFA Fellowship.

REFERENCES

A. D. Little Co. 1981. Human safety and environmental aspects of major

surfactants (Supplement). Report to the Soap and Detergent Association.

ADL Reference 84048.

ASTM (American Society for Testing and Materials). 2007. Standard Guide

for Conducting Acute Toxicity Tests on Material with Fishes,

Macroinvertebrates, and Amphibians. ASTM International, West Con-

shohocken.

ASTM (American Society for Testing and Materials). 2013. Standard Guide

for Conducting Laboratory Toxicity Tests with Freshwater Mussels.

ASTM International, West Conshohocken.

Archambault, J.M., C.M. Bergeron, W.G. Cope, R.J. Richardson, M.A.

Heilman, J.E. Corey III, M.D. Netherland, and R.J. Heise. 2015.

Sensitivity of freshwater molluscs to hydrilla-targeting herbicides:

Providing context for invasive aquatic weed control in diverse

ecosystems. Journal of Freshwater Ecology 30:335–348.

Arenzon, A., A.F. Pinto, P. Colombo, and M.T. Raya-Rodriguez. 2003.

Assessment of the freshwater annual fish Cynopoecilus melanotaenia as a

toxicity test organism using three reference substances. Environmental

Toxicology and Chemistry 22:2188-2190.

Augspurger, T. 2013. Use of Mussel Toxicity Data in Criteria, Standards, and

Risk Assessment. Presented at: SETAC North America 34th Annual

Meeting, 17-21 November 2013, Nashville. Society of Environmental

Toxicology and Chemistry.

Augspurger, T., A.E. Keller, M.C. Black, W.G. Cope, and F.J. Dwyer. 2003.

Water quality guidance for protection of freshwater mussels (Unionidae)

from ammonia exposure. Environmental Toxicology and Chemistry

22:2569-2575.

Barnhart, M.C. 2006. Buckets of muckets: A compact system for rearing

juvenile freshwater mussels. Aquaculture 254:227-233.

Belanger, S.E., D.M. Lee, J.W. Bowling, and E.M. LeBlanc. 2004. Responses

of periphyton and invertebrates to a tetradecyl-pentadecyl sulfate mixture

in stream mescosms. Environmental Toxicology and Chemistry 23:2202-

2213.

Berna, J.L., J. Ferrer, A. Moreno, D. Prats, and F. Ruiz Bevia. 1989. The fate

of LINEAR ALKYLBENZENE SULFONATE in the environment.

Tenside Surfactants Detergents 26:101-107.

Besser, J.M., D.L. Hardesty, I.E. Greer, and C.G. Ingersoll. 2009. Sensitivity

of freshwater snails to aquatic contaminants: Survival and growth of

endangered snail species and surrogates in 28-day exposures to copper,

ammonia and pentachlorophenol. Administrative Report CERC-8335-

FY07-20-10, submitted to USEPA. Duluth, MN. Office of Research and

Development.

Bringolf, R.B., W.G. Cope, C.B. Eads, P.R. Lazaro, M.C. Barnhart, and D.

Shea. 2007a. Acute and chronic toxicity of technical-grade pesticides to

glochidia and juveniles of freshwater mussels (Unionidae). Environmental

Toxicology and Chemistry 26:2086-2093.

Bringolf, R.B., W.G. Cope, C.B. Eads, P.R. Lazaro, M.C. Barnhart, and D.

Shea. 2007b. Acute and chronic toxicity of glyphosate compounds to

glochidia and juveniles of Lampsilis siliquoidea (Unionidae). Environ-

mental Toxicology and Chemistry 26:2094-2100.

Center for Biological Diversity. 2010. Petition to list 404 aquatic, riparian and

wetland species from the southeastern United States as threatened or

endangered under the Endangered Species Act. Center for Biological

Diversity, Tucson. Available at http://www.biologicaldiversity.org/

programs/biodiversity/1000_species/the_southeast_freshwater_

extinction_crisis/pdfs/SE_Petition.pdf (accessed September 2, 2014).

Chaturvedi, V., and A. Kumar. 2010. Toxicity of sodium dodecyl sulfate in

fishes and animals: a review. International Journal of Applied Biology and

Pharmaceutical Technology1:630-633.

Cowan-Ellsberry, C., S. Belanger, P. Dorn, S. Dyer, D. McAvoy, H.

Sanderson, D. Versteeg, D. Ferrer, and K. Stanton. 2014. Environmental

safety of the use of major surfactant classes in North America. Critical

Reviews in Environmental Science and Technology 44:1893-1993.

Conway, R.A., G.T. Waggy, M.H. Spiegel, and R.L. Berglund. 1983.

Environmental fate and effects of ethylene oxide. Environmental Science

and Technology 17:107-112.

Fogels, A., and J.B. Sprague. 1977. Comparative short-term tolerance of

Zebrafish, Flagfish, and Rainbow Trout to five poisons including potential

reference toxicants. Water Research 11:811-817.

Fox, K., M. Holt, M. Daniel, H. Buckland, and I. Guymer. 2000. Removal of

linear alkylbenzene sulfonate from a small Yorkshire stream: Contribution

to Greater Project #7. Science of the Total Environment 251:265-275.

Gibson, K.J. 2015. Acute toxicity testing on freshwater mussels (Bivalvia:

Unionidae) and freshwater snails (Gastropoda: Caenogastropoda). MS

Thesis. Troy University.

Graney, R.L., and J.P. Giesy, Jr. 1988. Alterations in the oxygen consumption,

condition index and centration of free amino acids in Corbicula fluminea

(Mollusca: Pelecypoda) exposed to sodium dodecyl sulfate. Environmen-

tal Toxicology and Chemistry 7:301-315.

Hemmer, M.J., M.G. Barron, and R.M. Greene. 2010. Comparative toxicity of

eight oil dispersant products on two Gulf of Mexico aquatic test species.

USEPA, Washington, D.C. Available at http://www.epa.gov/bpspill/

reports/Comparative ToxTest.Final.6.30.10.pdf (accessed December 16,

2014).

Hennes, E.C., and R.A. Rapaport. 1989. Calculation and analytical verification

of LINEAR ALKYLBENZENE SULFONATE concentrations in surface

waters, sediment and soil. Tenside, Surfactants, Detergents 26:141-147.

Holt, M.S., E. Matthus, and J. Waters. 1989. The concentrations and fate of

linear alkylbenzene sulfonate in sludge amended soils. Water Research

23:749-759.

Johnson, P.D., A.E. Bogan, K.M. Brown, N.M. Burkhead, J.R. Cordeiro, J.T.

Garner, P.D. Hartfield, D.A.W. Lepitzki, G.L. Mackie, E. Pip, T.A.

Tarpley, J.S. Tiemann, N.V. Whelan, E.E. Strong. 2013. Conservation

status of freshwater gastropods of Canada and the United States. Fisheries

38:247-282.

Kegley, S.E., B.R. Hill, S. Ome, and A.H. Choi. 2014. PAN Pesticide

Database. Pesticide Action Network, North America. Oakland, CA.

Available at http:www.pesticideinfo.org.

Keller, A.E. 1993. Acute toxicity of several pesticides, organic compounds,

34 GIBSON ET AL.

and a wastewater effluent to the freshwater mussel, Anodonta imbecillis,

Ceriodaphnia dubia, and Pimephales promelas. Bulletin of Environmen-

tal Contamination and Toxicology 51:696-702.

Keller, A.E., M. Lydy, and D.S. Ruessler. 2007. Unionid mussel sensitivity to

environmental contaminants. Pages 151-168 in J.L. Farris and J.H. van

Hassel, editors. Freshwater Bivalve Ecotoxicology. SETAC, Pensacola.

Khalladi, R., O. Benhabiles, F. Bentahar, and N. Moulai-Mostefa. 2009.

Surfactant remediation of diesel fuel polluted soil. Journal of Hazardous

Materials 164:1179-1184.

Lewis, M.A. 1991. Chronic and sublethal toxicities of surfactants to aquatic

animals: a review and risk assessment. Water Research 25:101-113.

Lewis, M.A., and V.T. Wee. 1983. Aquatic safety assessment for cationic

surfactants. Environmental Toxicology and Chemistry 2:105-118.

Lewis, P.A., and C.I. Weber. 1985. A study of the reliability of Daphnia acute

toxicity tests. Pages 73-86 in R.D. Cardnell, R. Purdy, and R.C. Bahner,

editors. Aquatic Toxicology and Hazard Assessment: 7th Symposium.

STP 854, ASTM. Philadelphia.

Misra, V., H. Lal, P.N. Viswanathan, and C.R.K. Murti. 1984. 45Ca uptake

from water by snails (Limnaea vulgaris) in control and detergent-polluted

samples. Ecotoxicology and Environmental Safety 8:97-99.

Neves, R.J., A.E. Bogan, J.D. Williams, S.A. Ahlstedt, and P.D. Hartfield.

1997. Status of aquatic mollusks in the southeastern United States: a

downward spiral of diversity. Pages 43-85 in G.W. Benz and D.E. Collins,

editors. Aquatic Fauna in Peril: The Southeastern Perspective. Southeast

Aquatic Research Institute, Special Publication, Chattanooga.

Newsome, C.S. 1982. Susceptibility of various fish species at different stages

of development to aquatic pollutants. Report EUR 7549, Community of

European Communities, Environmental Quality of Life, pp. 284-295.

Newton, T.J., and M.R. Bartsch. 2007. Lethal and sublethal effects of

ammonia to juvenile Lampsilis mussels (Unionidae) in sediment and

water-only exposures. Environmental Toxicology and Chemistry

26:2057-2065.

NIH (National Institute of Health). 2014. Household Product Database.

Available at http://householdproducts.nlm.nih.gov/cgi-bin/household/

brands?tbl¼chem&id¼78 (accessed November 20, 2014).

O Foighil, D., J. Li, T. Lee, P. Johnson, R. Evans, and J.B. Burch. 2011.

Conservation genetics of a critically endangered limpet genus and

rediscovery of an extinct species. PLoS ONE 6:1-9.

Ostroumov, S.A., and J. Widdows. 2006. Inhibition of mussel suspension

feeding by surfactants of three classes. Hydrobiologia 556:381-386.

Patrick, R., J. Cairns, Jr., and A. Scheier. 1968. The relative sensitivity of

diatoms, snails, and fish to twenty common constituents of industrial

wastes. The Progressive Fish-Culturist 30:137-140.

Reategui-Zirena, E.G., A. Whatley, F. Chu-Koo, and P.M. Stewart. 2013. Acute

toxicity testing of crude oil using Piaractus brachypomus, and Pimephales

promelas. International Journal of Environmental Protection 3:1-7.

Rebello, S., A.K. Asok, S. Mundayoor, and M.S. Jiha. 2014. Surfactants:

Toxicity, remediation and green surfactants. Environmental Chemistry

Letters 12:275-287.

Render, J. 1990. Effect of sodium dodecyl sulfate on polar lobe formation and

function in Ilyanassa obsolete embryos. Journal of Experimental Zoology

253:30-37.

Singer, M.M., and R.S. Tjeerdema. 1993. Fate and effects of the surfactant

sodium dodecyl sulfate. Review of Environmental Contamination and

Toxicology 133:95-149.

Susmi, T.S., S. Rebello, M.S. Jisha, and P.M. Sherief. 2010. Toxic effects of

sodium dodecyl sulphate on Grass Carp Ctenopharynogodon idella.

Fishery Technology 47:145.

Tarazona, J.V., and O. Nunez. 1987. Acute toxicity of synthetic detergents to

snails: Effects of sodium lauryl sulfate on Limnea peregra shells. Bulletin

of Environmental Contamination and Toxicology 16:1036-1040.

USEPA (U.S Environmental Protection Agency). 2002. Methods for

measuring the acute toxicity of effluent and receiving waters to freshwater

and marine organisms. Fourth Edition. EPA-821-R-02-012. U.S.

Environmental Protection Agency, Washington, D.C.

USEPA (U.S Environmental Protection Agency). 2009. National water quality

inventory - 2004. Report for Congress, Washington, D.C. Available at

http://water.epa.gov/lawsregs/guidance/cwa/305b/2004report_index.cfm

(accessed September 25, 2013).

USFWS (U.S Fish and Wildlife Service). 1993. Endangered status for eight

freshwater mussels and threatened status for three freshwater mussels in

the Mobile River drainage - Final Rule. Federal Register, 58 (60).

USFWS (U.S Fish and Wildlife Service). 1998. Endangered and threatened

wildlife and plants; endangered status for three aquatic snails, and

threatened status for three aquatic snails in the Mobile River basin of

Alabama - Final Rule. Federal Register, 63 (208).

van de Plassche, E.J., J.H.M. de Bruijn, R.R. Stephenson, S.J. Marshall, T.C.J.

Feijtel, and S.E. Belanger. 1999. Predicted no effect concentrations and

risk characterization of four surfactants: Linear alkyl benzene sulfonate,

alcohol ethoxylates, alcohol ethoxylated sulfates, and soap. Environmen-

tal Toxicology and Chemistry 18:2653-2663.

Verma, S.R., D. Mohan, and R.C. Dalela. 1978. Studies on the relative toxicity

of few synthetic detergents to a fish Macrones vittatus. Acta Hydro-

chimica et Hydrobiologica 6:121-128.

Verma, S.R., I.P. Tonk, and R.C. Dalela. 1981. Determination of the

maximum acceptable toxicant concentration (MATC) and the safe

concentration for certain aquatic pollutants. Acta Hydrochimica et

Hydrobiologica 9:247-254.

Villella, R.F., D. R. Smith, D. P. Lemarie. 2004. Estimating survival and

recruitment in a freshwater mussel population using mark-recapture

techniques. The American Midland Naturalist 151:114-133.

Wang, N., C.G. Ingersoll, I.E. Greer, D.K. Hardesty, C.D. Ivey, J.L. Kunz,

W.G. Brumbaugh, F.J. Dwyer, A.D. Roberts, T. Augspurger, C.M. Kane,

R.J. Neves, and M.C. Barnhart. 2007. Chronic toxicity of copper and

ammonia to juvenile freshwater mussels (Unionidae). Environmental

Toxicology and Chemistry 26:2048-2056.

Wang, N., C.G. Ingersoll, C.D. Ivey, D.K. Hardesty, T.W. May, T.

Augspurger, A.D. Roberts, E. van Genderen, and M.C. Barnhart. 2010.

Sensitivity of early life stages of freshwater mussels (Unionidae) to acute

and chronic toxicity of lead, cadmium, and zinc in water. Environmental

Toxicology and Chemistry 29:2053-2063.

Wang, N., C.A. Mebane, J.L. Kunz, C.G. Ingersoll, W.G. Brumbaugh, R.C.

Santore, J.W. Gorsuch, and W.R. Arnold. 2011. Influence of dissolved

organic carbon on toxicity of copper to a unionid mussel (Villosa iris) and

a cladoceran (Ceriodaphnia dubia) in acute and chronic water exposures.

Environmental Toxicology and Chemistry 30:2115-2125.

Wilcove, D.S., and L.L. Master. 2005. How many endangered species are there

in the United States? Frontiers in Ecology and Environment 3:414-420.

Williams, J.D., A.E. Bogan, and J.T. Garner. 2008. Freshwater Mussels of

Alabama & the Mobile Basin in Georgia, Mississippi & Tennessee. The

University of Alabama Press, Tuscaloosa.

35TOXICITY OF SODIUM DODECYL SULFATE TO FRESHWATER MOLLUSKS